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Latest Articles of Bio-Synthesis Inc.

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    Natural RNA tags added to RNA of interest allow detection, characterization, labeling and affinity purification of RNA target molecules. For example, small aptamer RNAs that bind with high affinity and specificity to Sephadex beads or streptavidin allow purification of intact RNA-protein complexes. Furthermore, RNA tags inserted into selected locations in genes encoding RNA components enable their detection. The CRISPR Cas system can be used together with synthetic RNA mimics to incorporate RNA tags into specific genomic locations to allow labeling.

    Recently a two-color CRISPR labeling system was developed by Wang and coworkers. Adding selective tags to a target-specific single guide RNA allows recruiting CRISPR-Cas systems for controlling gene expression.    

    Synthetic tagged RNA constructs allow purification of target RNAs and RNA complexes including miRNA and lncRNA as well as other molecules attached to the RNAs using pull-down techniques. RNA tags are unique tools for studying a variety of target RNAs including RNA transcripts, miRNAs, ncRNAs, lncRNAs, splicing event as well as others.

    Synthetic RNA can be chemically tagged or modified through the incorporation of modified ribonucleotides, such as BNAs, functional moieties such as biotin, fluorescent dyes, or natural or synthetic aptamers, for example, S1, D8, MS2 hairpin loops. Other types of RNA or constructs can be added as well.

    RNA affinity tags allow rapid enrichment of RNA binding protein (RNP) complexes from cellular lysates using mild conditions. This approach allows purification of RNP complexes under native conditions.

    Also, RNA tags can be used for the isolation of precursor or other RNA molecules of interest including RNPs.

    Known methods for the tagging of RNAs are: 

    (1)   Chemical tagging during in vitro transcription. 

    (2)   Incorporation of a well-characterized protein-binding RNA sequence
            during in vitro or in vivo transcription.

    (3)   Hybridization of affinity-tagged oligonucleotides that can also be biotinylated or
            modified, for example with BNAs.

    (4)   Incorporation of an artificially selected RNA motif during in vivo or
            in vitro transcription. 

    However, each of these methods has advantages and disadvantages.

    A variety of other RNA based applications are possible as well. Combining two or more hairpins within one RNA molecule is also possible.  

    Table 1:   Common RNA Tags

    Tag

    Sequence Info

    Notes

    Sephadex Tag

    D8 Sephadex RNA motif is recognized by Sephadex.

     

    5′ UCCGAGUAAU UUACGUUUUG  

       AUACGGUUGC GGA 3′

    The D8Sephadex-binding RNA minimal motif has 33 nucleotides. The indicated minimal structural motif has been discovered. The D8 tag was shown to bind specifically to Sephadex G-100 (Pharmacia).

    The D8 tag does not bind to other similar matrices such as Sepharose or Seph-acryl. binding of the tag to Sephadex can be efficiently competed with dextran B512.

    Steptavidin Tag

    S1 Streptavidin RNA motif recognized by streptavidin.

     

    5′ ACCGACCAGA AUCAUGCAAG

       UGCGUAAGAU AGUCGCGGGC

       CGGG 3′

     

    Kd of ~70 nM.

    The S1 streptavidin-binding RNA motif has 44 nucleotides originally selected to bind to streptavidin in either streptavidin–agarose bead assays or polyacrylamide gel electrophoretic mobility shift assays.

    Bound RNA tags can thus be released from streptavidin under otherwise native binding conditions by the inclusion of biotin in the binding buffer.

    MS2 Tag or

    MS2-TRAP

    Tagged RNA affinity purification.

    RNA is tagged with MS2 RNA hairpins and a fusion protein recognizing the MS2 RNA hairpins, MS2-GST is used.

    Identification of miRNAs associated with a target transcript in the cellular context.

    Identification of microRNAs that associate with a long intergenic (li)ncRNA.

    BoxB sequence

     = Boxb Rna

    NGTTCACCTCTAACCGGGTGAGCC

     

    Is recognized by the bacteriophage protein λ N:

     

    N Peptide: 

    ESKGTAKSRYKARRAELIAERR 

    BoxB can be used to tether proteins to RNAs, for example to mRNAs.

    PP7 Hairpin

    Binds to PP7 coat protein.

    Different RNA hairpins are rcognized by the coat proteins of different single-stranded RNA phages.

     

    Proposed and solved Structures of Hairpins


    Figure 1: Minimal binding motif and consensus structures of the Sephadex-affinity tag.

    Figure 2: Minimal binding motif and consensus structures of the streptavidin-affinity tag. (X indicates nonconserved nucleotides).

    Figure 3: Different views of the structural model of a MS2-RNA hairpin (G-5) complex. (Source: PDB 2C51). 

    Figure 4: Different views of the structural model of the MS2-RNA hairpin (G-5). (Source: PDB 2C51).

    pdb|1NYB|A Chain A, Solution Structure Of The Bacteriophage Phi21
               N Peptide-Boxb Rna Complex  ESKGTAKSRYKARRAELIAERR
    pdb|1NYB|B Chain B, Solution Structure Of The Bacteriophage Phi21
               N Peptide-Boxb Rna Complex  NGTTCACCTCTAACCGGGTGAGCC

    Figure 5: Two views of the structural models from the solution structure of a 22-amino-acid peptide from the amino-terminal domain of the bacteriophage φ21 N protein in complex with its cognate 24-mer boxB RNA hairpin solved with NMR.

    The nut (N utilization) site of bacteriophage lambda consists of two genetically defined elements, boxA and boxB. boxB forms an RNA hairpin and its 5 bp stem and 5 nt loop are recognized by the N peptide. The N peptide binds as an α-helix and interacts predominately with the major groove side of the 5′ half of the boxB RNA stem-loop. The φ21 boxB loop (CUAACC) has a structure typical of the “U-turn” motif.

    Figure 6: Two views of the PP7 coat protein dimer in complex with RNA hairpin. The RNA hairpin binds across the β-sheet surface of the coat protein dimer.

    The combined use of the MS2 hairpin with the PP7 hairpin allows detection of RNA in live cells if a chimeric protein consitent of the phage protein, a nuclear localization signal, and a fluorescent molecules is used. The MS2/PP7 appraoch has been used for the study of movement and localization of RNA as well the formation of RNA at the transcription site.    

    Reference

    Baron-Benhamou J, Gehring NH, Kulozik AE, Hentze MW.; Using the lambdaN peptide to tether proteins to RNAs. Methods Mol Biol. 2004;257:135-54. https://www.ncbi.nlm.nih.gov/pubmed/14770003
    Chao, J. A., Patskovsky, Y., Almo, S. C., & Singer, R. H. (2008). Structural basis for the coevolution of a viral RNA–protein complexNature Structural & Molecular Biology15(1), 103–105. 
    Cilley CD, Williamson JR.; Structural mimicry in the phage phi21 N peptide-boxB RNA complex. RNA. 2003 Jun;9(6):663-76. https://www.ncbi.nlm.nih.gov/pubmed/12756325

    Gesnel, M.-C., Del Gatto-Konczak, F., & Breathnach, R. (2009). Combined Use of MS2 and PP7 Coat Fusions Shows that TIA-1 Dominates hnRNP A1 for K-SAM Exon Splicing Control. Journal of Biomedicine and Biotechnology, 2009, 104853. http://doi.org/10.1155/2009/104853.

    Lenstra, T. L., & Larson, D. R. (2016). Single-Molecule mRNA Detection in Live YeastCurrent Protocols in Molecular Biology / Edited by Frederick M. Ausubel ... [et Al.]113, 14.24.1–14.24.15. http://doi.org/10.1002/0471142727.mb1424s113.

    Lo, A., & Qi, L. (2017). Genetic and epigenetic control of gene expression by CRISPR–Cas systems . F1000Research6, F1000 Faculty Rev–747. 

    Francis Lim and David S. Peabody; RNA recognition site of PP7 coat protein.Nucleic Acids Res. 2002 Oct 1; 30(19): 4138–4144. PMCID: PMC140551



    Marchese, D., de Groot, N. S., Lorenzo Gotor, N., Livi, C. M., & Tartaglia, G. G. (2016). Advances in the characterization of RNA
    binding proteins. Wiley Interdisciplinary Reviews. RNA, 7(6), 793–810. http://doi.org/10.1002/wrna.1378.

    Pascale Legault, Joyce Li, Jeremy Mogridge, Lewis E Kay, Jack Greenblatt; NMR Structure of the Bacteriophage λ N Peptide/boxB RNA Complex: Recognition of a GNRA Fold by an Arginine-Rich Motif. Cell, Volume 93, Issue 2, 1998, 289-299.https://doi.org/10.1016/S0092-8674(00)81579-2. (http://www.sciencedirect.com/science/article/pii/S0092867400815792)

    Srisawat, C., & Engelke, D. R. (2002). RNA affinity tags for purification of RNAs and ribonucleoprotein complexes. Methods (San Diego, Calif.), 26(2), 156–161. http://doi.org/10.1016/S1046-2023(02)00018-X

    Walker, S. C., Scott, F. H., Srisawat, C., & Engelke, D. R. (2008). RNA Affinity Tags for the Rapid Purification and Investigation of RNAs and RNA–Protein Complexes. Methods in Molecular Biology (Clifton, N.J.), 488, 23–40. http://doi.org/10.1007/978-1-60327-475-3_3

    Wang, S., Su, J.-H., Zhang, F., & Zhuang, X. (2016). An RNA-aptamer-based two-color CRISPR labeling system. Scientific Reports, 6, 26857. http://doi.org/10.1038/srep26857.

    Yoon, J.-H., Srikantan, S., & Gorospe, M. (2012). MS2-TRAP (MS2-tagged RNA affinity purification): Tagging RNA to identify associated miRNAs. Methods (San Diego, Calif.), 58(2), 81–87. http://doi.org/10.1016/j.ymeth.2012.07.004

    Yoon, J.-H., & Gorospe, M. (2016). Identification of mRNA-interacting factors by MS2-TRAP (MS2-tagged RNA affinity purification). Methods in Molecular Biology (Clifton, N.J.), 1421, 15–22. http://doi.org/10.1007/978-1-4939-3591-8_2

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    Bioinformatic algorithms allowing predicting of biomolecular folding for proteins, peptides, and RNAs, even though sometimes successful, have all their limitations. The reasons are as follows.

    •   RNA molecules in solution may adopt secondary structures that are only partially
         determined by thermodynamics since RNA molecules can undergo conformational
         changes during interaction with other RNAs, RNA binding proteins or RNA binding
         peptides. These interactions are very complex and difficult to model.

    •   Our knowledge of thermodynamic rules and parameters that govern folding
         patterns of RNAs are far from being complete.

    •   Most folding algorithms use approximations for scanning the landscape of possible
         secondary structures. Therefore the predicted structures are often just an
         estimate or approximation and chemical methods are needed to verify the
         deduced structures.

    •   Predicting pseudoknots and long-range and tertiary-structure interactions
         accurately is also quite difficult.

    •   Using experimental methods to accumulate additional experimental data
         should allow improvements in algorithms used in computational methods.    

    The mapping of RNA-protein or RNA-RNA interactions by protein pull-down or affinity pull-down methods allow studying RNA structures, as well as RNA-protein, and RNA-RNA interactions. Since RNA-binding proteins (RBPs) are key players in the post-transcriptional regulation of gene expression precise knowledge of their binding sites is critical for determining their molecular function and for understanding their roles in cell development and disease.

    NMR

    Nuclear magnetic resonance spectroscopy is a powerful tool for studying RNA structures in detail. NMR allows studying RNA molecules in a more natural state when dissolved in solution. The drawback is that a large preparation of highly purified and uniform RNA is needed, and it is limited to solving small structures. 
    https://www.ncbi.nlm.nih.gov/pubmed/14523911,  https://www.nature.com/articles/ncomms8024.


    RIP

    RNA immunoprecipitation (RIP) uses antibodies to pull down RNA bound to a targeted protein. RIP allows detection of individual proteins associated with specific nucleic acid regions on RNA molecules. Live cells are treated with formaldehyde to generate protein-RNA cross-links between molecules that are nearby. Cross-linked RNA connected to the targeted protein is isolated using immunoprecipitation of the protein. Recovery and quantitative analysis of the immunoprecipitated RNAs are achieved by reversal of the formaldehyde cross-linking which now permits reverse transcription using PCR. RIP is similar to chromatin immunoprecipitation (ChiP). Unfortunately, the technique cannot differentiate between direct and indirectly bound RNA. A drawback of this method is that the generation of false positives from interaction occurring after cell lysis is also possible. 
    https://www.ncbi.nlm.nih.gov/pubmed/27659976


    CLIP

    Cross-linking and immunoprecipitation (CLIP) allows mapping of transcriptome-wide binding sites of RNA-binding proteins. RNA-protein complexes are covalently cross-linked and purified from intact tissue cells. CLIP improved the specificity of RIP. It allows the removal of weekly bound RNA when stringent washing steps are used. RNAs that remain can be reversed transcribed, and PCR amplification allows sequencing with next-generation sequencing. For CLIP to work, reverse transcription needs to proceed from a universal 3′ ligated adapter to a universal 5′ ligated adapter. Both adapters are required for PCR amplification.

    All CLIP protocols use RNA-protein cross-linking and immunoprecipitation targeting a protein of interest.  Ultraviolet light creates the cross-links between RNA and proteins in vivo. The RNA is then isolated and reverse transcribed into cDNA. The cDNA can be used on several platforms to identify and quantify interacting RNAs. Several refinements and specializations of this central CLIP principle exist: CLIP-seq, PAR-CLIP, and iCLIP are three of the most common.

    http://epigenie.com/quick-review-crosslinking-immunoprecipitation-clip-methods/
    https://www.ncbi.nlm.nih.gov/pubmed/3071669
    http://science.sciencemag.org/content/302/5648/1212.long
    https://www.ncbi.nlm.nih.gov/pubmed/16314267?dopt=Abstract
    https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4492529/https://en.wikipedia.org/wiki/CLIP


    iCLIP

    The individual-nucleotide resolution CLIP (iCLIP) protocol was developed to allow recovery of truncated cDNAs lost in CLIP. iCLIP enables PCR amplification of truncated cDNAs and identifies protein–RNA crosslink sites with single nucleotide resolution. iCLIP identifies protein-RNA cross-links on a genome-wide scale. An intramolecular cDNA circularization step enables analysis of cDNAs truncated at the protein-RNA crosslink sites with high resolution and specificity. A 3’ exonuclease degrades protein-bound RNA. The enzyme digests the isolated RNA but stops at the cross-linked protein. An adapter is then ligated to the remaining RNA.

    For iCLIP, cells are irradiated with UV-C light on ice. Covalent bonds are formed between proteins and RNA. The cross-linking reaction is followed by partial RNase digestion and immunoprecipitation with protein-specific antibodies. Libraries are preparation and visualized by dephosphorylation of RNA. Next, a 3′ end adapter is ligated to the RNA, and the 5′ end is radioactively labeled. Complexes are separated by SDS–PAGE and isolated from a nitrocellulose membrane. Proteins are digested by proteinase K, and reverse transcription (RT) is performed truncating at the remaining polypeptide. An RT primer introduces two cleavable adapter regions and barcode sequences. Free RT primers are removed by size selection, and circularization of the cDNA is carried out. Linearization generates suitable templates for PCR amplification.Finally, high-throughput sequencing generates sequencing reads in which a barcode sequence is immediately followed by the last nucleotide of the cDNA.
    https://www.ncbi.nlm.nih.gov/pubmed/24184352 
    https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3988997/
    http://epigenie.com/quick-review-crosslinking-immunoprecipitation-clip-methods/


    eCLIP

    Enhanced CLIP improves library preparation and circular steps of iCLIP. eCLIP simplifies the generation of paired IgG and size-matched input controls. Also, eCLIP improves specificity in the discovery of authentic binding sites. After dephosphorylation of RNA fragments, an “inline barcoded” RNA adapter is ligated to the 3′ end. Following protein gel electrophoresis and nitrocellulose membrane transfer, a region of 75 kDa (~220 nt of RNA) above the protein size is excised and proteinase K treated to isolate RNA. The RNA is further prepared into paired-end high-throughput sequencing libraries and sequenced.
    https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4887338/

    PAR-CLIP

    Photo-activable ribonucleoside-enhanced cross-linking and immunoprecipitation (PAR-CLIP) is a method that allows identification of binding sites of RNA-binding proteins (RBPs). PAR-CLIP can be used for the identification of RNA-protein binding sites in transcriptomes. For the method to work, photoreactive ribonucleoside analogs are incorporated into nascent RNA transcripts in living cells. The use of ultraviolet (UV) light of 365 nm cross-links photoreactive nucleoside-labeled cellular RNAs to RNA-binding proteins.

    First, photo-reactive thioribonucleosides are incorporated into nascent transcripts. Cross-linking is achieved by irritating the cells with ultraviolet long-wavelengths greater than 310 nm (365 nm is usually used).  Immunoprecipitation is used for the purification of cross-linked RNA–RBP complexes which are further purified by SDS-PAGE. The recovered RNA is converted into a cDNA library and is sequenced using next-generation sequencing. Multiple sequencing platforms can be used.

    Reverse transcription of cross-linked RNA with incorporated photoactivable thioribonucleosides, followed by PCR amplification, results in a characteristic mutation (T-to-C when using 4SU and G-to-A when using 6SG) that is used to identify the RNA recognition elements. 
    https://www.ncbi.nlm.nih.gov/pubmed/29236251


    DMS Treatment

    Dimethylsulfate (DMS, C2H6O4S, 126.13 g/mol) treatment modifies RNA by adding a methyl group to any unpaired or loosely structured adenosine (A) and cytidine (C) in oligonucleotides. DMS mapping is one of the oldest chemical RNA mapping methods. After methylation, the bases can no longer form base pairs and will cause cDNA transcripts to terminate early allowing identification of the presence of unpaired bases. The following use of next-generation-sequencing (DMS-seq/Structure-seq) increases the power of the technique and allows quantitative mapping of base modifications. Targeted Structure-seq further improves the specificity of the technique by using primers targeting the length of a specific RNA of interest without having to sequence the whole transcriptome. 
    https://www.ncbi.nlm.nih.gov/pmc/articles/PMC2701642/


    PARS

    Parallel analysis of RNA structure (PARS) combines classical RNA foot-printing methods with next-generation sequencing. Here, poly(A) selected RNA is folded in vitro and incubated with either RNase V1 or S1 nuclease to probe for double- and single-stranded regions. RNase V1 and S1 nuclease cleavage results in a 5′P leaving group. Nest, the RNA is fragmented. Enzymatic cleavage products contain 5′P, but fragmentation and degradation products have 5′OH groups. Only true structure-probing sites can be ligated to adaptors and reverse transcribed. Sequencing the resulting cDNA library using high-throughput sequencing and mapping the resulting reads to the genome allow identification of double- or single-stranded regions in the transcriptome. Different PARS scores indicate that a base is either double-stranded (positive score) or single-stranded (negative score).


    PARIS

    Psoralen analysis of RNA interactions and structure (PARIS) is a psoraleb based cross-linking method that works by fixing base pairs of double-stranded RNA (dsRNA) of cells in vivo using the specific cross-linker 4′-aminomethyl-trioxsalen (AMT). PARIS analysis directly determines transcriptome-wide base pairing interactions. PARIS combines in vivo cross-linking, 2D gel purification, proximity ligation, and high-throughput sequencing to allow high-throughput and near-base pair resolution determination of the RNA structures and interactions in living cells. After cross-linking sample are treated with proteinase and the RNA is partially degradated. The resulting sequencing reads represent all native dsRNAs in the organism. These can be mapped to infer their structure.
    https://www.ncbi.nlm.nih.gov/pubmed/29130190


    Psoralen conjugates oligonucleotides

    Psoralen conjugates oligonucleotides can also be used as antisense oligonucleotides (ASOs). The use of triple helix-forming oligonucleotides linked to psoralen (pso-TFO) allows introduction of DNA interstrand cross-links at specific sites in the genome of living mammalian cells. 
    Co-introduction of duplex DNA with target region homology results in precise knock-in of the donor at frequencies 2–3 orders.

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    The development of next-generation sequencing (NGS) methods revolutionized the study of genomic transcriptionally active regions. NGS now makes it possible to catalog transcripts from coding sequences and also allows the discovery of RNA species that are not templates for protein synthesis.

    Several classes of non-coding RNAs (ncRNAs), including microRNAs (miRNAs) and long non-coding RNAs (ncRNAs), are already known to play diverse roles in post-transcriptional regulation of mRNA stability and epigenetic control of chromatin activities.

    Recently a new class of RNAs called enhancer RNAs or eRNAs has been identified {Kim et al. 2010}. However, it remains an open question whether eRNAs are just transcriptional noise or have a relevant biologically function. As usually, to clarify this, more research is needed.

    Figure 1: Overview of a typical gene transcribed by RNA polymerase II.

    A typical gene transcribed by RNA polymerase II has a promotor that extends upstream from the site where transcription is initiated (Figure 1).  (See Lewin, Benjamin; Genes VII chapter 20, 2000). The promotor contains several short sequence elements that bind transcription factors, up to 10 base pairs (bp) in length, and promotors are dispersed over a sequence region that can be greater than 200 bp. Enhancers contain a more closely packed array of elements that also bind transcription factors. Enhancer regions may be located at a distance of several kilobases (kb). The DNA duplex may be coiled or rearranged such that transcription factors at the promotor and the enhancer interact to form a large protein-DNA complex.

    What are eRNAs?

    Enhancer RNA or eRNA are RNAs transcribed by RNA polymerase II (RNAPII) from the domain of transcription enhancers. eRNAs are a class of short non-coding RNAs, 50 to 2,000 nucleotides in length, transcribed from DNA enhancer regions. eRNAs stimulate gene expression, but the precise mechanisms how they function remains unclear. Enhancers are intergenic DNA elements that regulate transcription of target genes in response to signaling pathways by interacting with promotors over large genomic distances. Enhancers contain binding sites for transcription factors that promote RNA polymerase II (RNAPII) recruitment and transcription activation. Also, enhancers carry unique epigenetic marks that distinguish them from promoters. Additionally, these regulatory elements have an open chromatin conformation that increases accessibility to transcription factors and RNAPII. Enhancers are bi-directionally transcribed into eRNAs {Kim et al. 2010}.

    Rahman et al. recently reported that eRNAs are localized exclusively in the nucleus and the induction of eRNAs occurs with similar kinetics as that of target mRNAs. eRNAs are mostly nascent at enhancers however their steady-state levels remain lower than those of their cognate mRNAs at the single-allele level. eRNAs are rarely co-expressed with their target loci. It appears that active gene transcription does not require the continuous transcription of eRNAs or their accumulation at enhancers. 

    Genome-wide sequencing methods allowed studying stimulus-dependent enhancer functions in tissues cells. Kim et al. in 2010 found that the level of eRNA expression at neuronal enhancers positively correlates with the level of mRNA synthesis in nearby genes. Their findings suggest that eRNA synthesis occurs specifically at enhancers that actively promote mRNA synthesis and the mechanism of enhancer activation involves RNAPII binding and eRNA synthesis.

    Levels of eRNA expression correlate with mRNA levels of the corresponding enhancers target gene. eRNAs can be identified by a specific chromatin signature: H3K4me1 and H3K27ac. Furthermore, enhancers are often transcribed and bound by RNA polymerase II. eRNAs appear to be rarely spliced, are not polyadenylated, and are often transcribed in both directions. So far, the mechanisms by which eRNAs facilitate enhancer function remain unclear. More research to clarify this is needed.

    Kim et al. suggested that eRNA synthesis is required to establish and maintain a chromatin landscape at enhancers needed for enhancer function. It is also possible that the eRNA transcripts are functionally important by themselves.

    Enhancers are classified as cis-regulatory genetic elements that controll temporal and cell-type specific patterns of gene expression. Active enhancers have been found to generate bi-directional non-coding RNA transcripts called enhancer eRNAs. Enhancers contain bidirectional elements that allow assisting initiation and the activity of a promotor is increased by the presence of an enhancer. The enhancer is located distinct from the promotor and the position of the enhancer relative to the promotor can vary substantially. An enhancer can stimulate any promotor placed in its vicinity. Enhancers often show redundancy in function. DNA must be able to form a loop structure if proteins bound at an enhancer several kb distant from a promotor need to interact directly with proteins bound in the vicinity of the starting point such that the enhancer and promotor are closely located to each other. Enhancers may function by bringing proteins into the vicinity of the promotor. A model for eRNAs and transcriptional activation of a typical gene is illustrated in figure 2.

    Figure 2: Model for eRNAs, enhancer derived RNAs, and transcriptional activation of a gene (Feng Liu, 2017). The structure of a typical gene is illustrated. The gene is associated with two cis-regulatory elements, (i) a promotor, located in proximity, and (ii) an enhancer, located distal to the transcription start site of the gene. In general, when its enhancer is inactivae the gene is turned “off” (A). When an enhancer is activated by transcription factors, it can loop towards the promotor and turn “on” the transcription of the gene (B). Both, enhancers and promotors are classified as non-coding elements.



    A more detailed model can be reviewed in the paper published by Kim et al. in 2015 in which the loop structure is shown together with the transcription machinery.

    To find and varify the presence of eRNAs, Espinosa suggested in 2015 to design a couple of CRISPRs, one to delete the lncRNA "promotor" and a second one to induce a pol(A) cassette early in the path of RNAPII. If the first genome edit affects expression but the second one doesn't it is an "e" or eRNA. 



    Reference

    Espinosa JM; Revisiting lncRNAs: How Do You Know Yours Is Not an eRNA? Mol Cell. 2016 Apr 7;62(1):1-2. doi: 10.1016/j.molcel.2016.03.022.

    Giles et al. 2015. ncRNA function in chromatin organization., in Epigenetic Gene Expression and Regulation}.

    Kim, T.-K., Hemberg, M., Gray, J. M., Costa, A. M., Bear, D. M., Wu, J., … Greenberg, M. E. (2010).
    Widespread transcription at neuronal activity-regulated enhancers. Nature, 465(7295), 182–187. http://doi.org/10.1038/nature09033. https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE21161.

    Kim, T-K., Hemberg, M., and Gray J.M.; 2015. Enhancer RNAs: A class of long noncoding RNAs synthesized at enhancers. Cold Spring Harb. Perspect 2015,7:a018622, 1-3. 

    Lewin, Benjamin; Genes VII chapter 20, 2000

    Feng Liu; Enhancer-derived RNA: A Primer. Genomics Proteomics Bioinformatics 15 (2017) 196-200.

    Rahman, S., Zorca, C. E., Traboulsi, T., Noutahi, E., Krause, M. R., Mader, S., & Zenklusen, D. (2017).
    Single-cell profiling reveals that eRNA accumulation at enhancer–promoter loops is not required to sustain transcription. Nucleic Acids Research, 45(6), 3017–3030. http://doi.org/10.1093/nar/gkw1220.

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    BNAs dramatically improve the specificity of CRISPR-Cas9. 

    The addition of BNAs (2′,4′-BNANC[N-Me]) to CRISPR-RNAs (crRNAs) dramatically improves the accuracy in CRISPR-based gene-editing. In this approach, natural crRNA molecules are replaced with synthetic crRNAs containing bridged nucleic acids, or BNAs.

    BNA-T (2′,4′-BNANC[N-Me]-T)

    Off-target effects or off-target cutting and the generation of additional mutations remain a significant barrier for the use of Cas9-based gene editing. Therefore a number of approaches have been investigated and developed to improve the specificity of Cas9 and to minimize or abolish off-target effects. 

    Cromwell et al. in 2018 now report that the incorporation of bridged nucleic acids, BNAs, at specific positions within crRNAs significantly improves Cas9 DNA cleavage specificity in vitro and cells with close to no off-target cleavage. The researcher reasoned that modified crRNAs enhance specificity by impairing the formation of the stable “zipped” conformation during hybridization to off-target sequences.

    These results indicate that BNA modified crRNAs improve the specificity of the CRISPR-Cas9 system and illustrate the power of recently developed synthetic nucleic acid technologies to solve problems in enzyme specificity as well.

    The research was published in "Nature Communications" in a paper entitled “Incorporation of Bridged Nucleic Acids into CRISPR RNAs Improves Cas9 Endonuclease Specificity.”

    The CRISPR (Clustered Regularly Interspaced Short Palindromic Repeat)-Cas9 complex is a component of prokaryotic immune systems that is now a widely used tool for genome editing technologies. CRISPR-Cas9 can be used for the generation of gene knock-outs and knock-ins in a variety of organisms, functional genomics studies, epigenetic screens, and proof-of-principle studies that enable the correction of genetic diseases in mammals.

    Reference

    Cromwell, Christopher R., Sung, Keewon, Park, Jinho, Krysler, Amanda R., Jovel, Juan, Kim, Seong Keun, and Hubbard, Basil P.; 2018. Incorporation of bridged nucleic acids into CRISPR RNAs improves Cas9 endonuclease specificity. Nature Communications 9, 1448.


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    Horseradish Peroxidase, or HRP, allows the design and synthesis of molecular probes such as in situ hybridization probes to increase signal amplification. HRP, when conjugated to hybridization probes, increases signal amplification, for example, in Catalyzed Reporter Deposition-Fluorescent In Situ Hybridization (CARD-FISH) applications.

    Historically, horseradish peroxidase is used as a reporter enzyme in diagnostics and histochemistry applications. Horseradish Peroxidase can be conjugated to 
    oligonucleotides, peptides, proteins, or other molecules, for example, via a maleimide attachment to form a thioester bond.


    Horseradish Peroxidase is an enzymatically active glycoprotein. HRP is a globular molecule with an α-helical secondary structure and one short β-sheet region (Gajhede et al. 1997). 

    The HRP molecule is separated into a distal and a proximal region by an iron protoporphyrin IX cofactor. The iron protoporphyrin IX cofactor is known as the heme group. Heme is typically linked to HRP by a coordinate bond of the heme iron with a conserved His170 residue. In the presence of H2O2, HRP catalyzes an oxidation reaction that converts reduced, non-fluorescent or non-colorimetric substrates into oxidized, fluorescent or colorimetric molecules. Furthermore, HRP C1A contains nine Asn-X-Ser/Thr-X motives (X being any amino acid but Pro) which are potential N-glycosylation sites.

    For conjugation reactions, surface lysine residues allow directed enzyme immobilization or conjugation to target compounds via covalent linkages. On the surface of HRP C1A, three out of six lysine residues were found accessible to chemical modifications. These are 
    Lys174, Lys232, and Lys241 (O’Brien et al. 2001). 


    Reference

    Ferrari, B. C., Tujula, N., Stoner, K., & Kjelleberg, S. (2006). Catalyzed Reporter Deposition-Fluorescence In Situ Hybridization Allows for Enrichment-Independent Detection of Microcolony-Forming Soil Bacteria. Applied and Environmental Microbiology, 72(1), 918–922. [PubMed]

    Gajhede M, Schuller DJ, Henriksen A, Smith AT, Poulos TL. Crystal structure of horseradish peroxidase C at 2.15 A resolution. Nat Struct Biol. 1997;4:1032–8. [
    PubMed]

    Krainer, F. W., & Glieder, A. (2015). An updated view on horseradish peroxidases: recombinant production and biotechnological applications. Applied Microbiology and Biotechnology, 99, 1611–1625. 

    [PubMed]

    O’Brien AM, O’Fágáin C, Nielsen PF, Welinder KG. Location of crosslinks in chemically stabilized horseradish peroxidase: implications for the design of crosslinks. Biotechnol Bioeng. 2001;76:277–84. [
    PubMed]


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    To increase solubility and effectiveness of Paclitaxel, the conjugation of the drug to gold nanoparticles via DNA linkers is possible.  According to Zhang et al., resulting conjugates usually show a 50-fold increase in solubility in aqueous buffers in comparison to the unconjugated drug. For visualization of the conjugation reaction and the effect of the drug conjugate in cells, the DNA linker can be labeled with fluorophores as well (Zhang et al. 2011). The final nano-gold DNA Paclitaxel conjugate represents a multimodal drug delivery system with

    (i)  enhanced solubility of the drug in aqueous systems, and serum-containing
         cell culture media,

    (ii) increased drug efficacy in Paclitaxel-resistant cells, and

    (iii) a useful way of tracking the delivery of the drug. 

    However, other types of Paclitaxel conjugates are also possible.



    Taxol

    .

    Paclitaxel, Taxol A,  (Source: Drugbank)

    (2AR-(2aalpha,4beta, 4abeta,6beta, 9alpha(alpha r*,betas*), 11alpha,12alpha,12balpha))-beta-(benzoylamino)-alpha- hydroxy-benzenepropanoic acid 6,12b-bis(acetyloxy)-12-(benzoyl-oxy)-2a,3,4,4a,5,6,9,10,11,12,12a,12b-dodecahydro-4,11-dihydroxy-4a,8,13,13-tetramethyl-5-oxo-7,11-methano-1H-cyclodeca(3,4)benz(1,2-b)oxet-9-yl ester. 

    C47H51NO14.

    Mw: Average: 853.9061 
    ; Monoisotopic: 853.330955345


    “Paclitaxel is a mitotic inhibitor used in cancer chemotherapy. It was discovered in a US National Cancer Institute program at the Research Triangle Institute in 1967 when Monroe E. Wall and Mansukh C. Wani isolated it from the bark of the Pacific yew tree, Taxus brevifolia and named it taxol. Later it was discovered that endophytic fungi in the bark synthesize paclitaxel. When it was developed commercially by Bristol-Myers Squibb (BMS), the generic name was changed to paclitaxel and the BMS compound is sold under the trademark Taxol. In this formulation, paclitaxel is dissolved in Kolliphor EL and ethanol, as a delivery agent. A newer formulation, in which paclitaxel is bound to albumin, is sold under the trademark Abraxane”.  [Wikipedia, Drugbank]
     

    Taxol binds to microtubules in the cell and slows down cell division and growth. It does so by stabilizing microtubules thereby blocking the segregation of chromosomes resulting in cell death or cellular apoptosis. However, formulation of Taxol into a delivery system acceptable for human use proved difficult. According to the NIH, Taxol is the best-selling drug ever manufactured.

    Different illustrations of the tubulin structure with highlighted GTP (non-exchangeable) bound to α-tubulin, GDP (exchangeable) bound to β-tubulin, and Taxol.




     

    Due to its inherent insolubility in aqueous media, Paclitaxel can be limited in effectiveness. However, the covalent attachment to gold nanoparticles via DNA linkers results in conjugates highly soluble in aqueous buffers. Zhang et al. in 2011 reported a 50 fold increase in solubility of the conjugates when compared to the unconjugated drug.

    The research group recommends the attachment of DNA-nanoparticle conjugates to drugs as a strategy for the enhancement of a wide variety of therapeutic agents that need to be formulated in aqueous media.  


    Reference

    Hermanson, Grg T.; Bioconjugate Techniques.

    Kim CK, Ghosh P, Rotello VM.;  Multimodal drug delivery using gold nanoparticles. Nanoscale. 2009 Oct;1(1):61-7. doi: 10.1039/b9nr00112c. Epub 2009 Sep 4.

    Zhang, X.-Q., Xu, X., Lam, R., Giljohann, D., Ho, D., & Mirkin, C. A. (2011). A Strategy for Increasing Drug Solubility and Efficacy through Covalent Attachment to Polyvalent DNA-Nanoparticle Conjugates. ACS Nano, 5(9), 6962–6970. http://doi.org/10.1021/nn201446c

    ---...---


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    To overcome disadvantages of drugs with poor water solubility different strategies such as the conjugation of the drug to small molecules, carbohydrates, oligonucleotides, peptides, or other molecules have been developed by many research groups from all over the world in recent decades.

    The solubility of the drug Paclitaxel in aqueous media can be increase by conjugating Paclitaxel to aptamers, oligonucleotides, or DNA linked gold nanoparticles.

    A good example is the conjugation of functionalized Paclitaxel to gold nanoparticles via a DNA linker. In this approach Zhang et al. used succinic anhydride to add a functional extender group to the drug that now allows conjugation to oligonucleotides containing an amino group on one of it's terminal ends followed by conjugating this conjugate to nano-gold particles. First, Paclitaxel is reacted with succinic anhydride to generated Paclitaxel carboxylic acid which now allows conjugation to an amino group on a DNA oligonucleotide.

    The following steps were needed for the production of fluorescent Paclitaxel-DNA-gold nanoparticles.


    (1)    Synthesize propyl thiol-(3’-end) DNA oligomers with a terminal amino group
             (5’-end) needed for covalent attachment of Paclitaxel.

    (2)    Modify Paclitaxel with succinic anhydride to create Paclitaxel carboxylic acid.

    (3)    Covalently attach Paclitaxel carboxylic acid to the modified oligonucleotide
             using EDC/Sulf-NHS.

    (4)    Purify and QC the derivative (PTX-DNA) using MALDI-MS.

    (5)    Immobilize the derivative on citrate-stabilized gold nanoparticles (AuNPs).

    (6)    Remove excess PTX-DNA.

    (7)    Resuspend PTX-DNA-AuNPs.

    For the synthesis of the Paclitaxel-fluorescein-DNA conjugate (Zhang et al. 2011) succinic anhydride was used to modify Paclitaxel by the reaction with succinic anhydride to create Paclitaxel carboxylic acid. 




    A modified DNA oligonucleotide containing a amino group at the 5’-end, an internal fluorophore, and a thiol group at the 3'-end was covalently attached to Paclitaxel carboxylic acid to yield the Paclitaxel-DNA (PTX-DNA) compound. (Structural models are not to scale).


    Immobilization of PTX-DNA on citrate-stabilized gold nanoparticles (AuNPs) yielded the Paclitaxel-DNA gold nanoparticle conjugate (PTX-DNA@AuNPs). HPLC is used for the purification of intermediate products and MALDI-TOF-MS is used for QA/QC. 



    Zhang et al. showed that covalently attached hydrophobic Paclitaxel onto gold nanoparticles via a DNA linker resulted in enhanced hydrophilicity and stability of the conjugate. Cellular internalization, delivery, and distribution of the drug conjugate was monitored in human breast adenocarcinoma cells and uterine sarcoma cells with the help of confocal fluorescence microscopy. Furthermore, the cell-killing activity of paclitaxel was enhanced in vitro against several cancer cell lines when delivered as the gold nanoparticle conjugate.

    The authors of the referenced papers suggested that this approch and other similar approaches can be used for other biological useful compounds such as peptides, small interfering RNA (siRNA), gadolinium complexes, antibodies, as well as aptamers.

    To conclude, these types of conjugates represent a powerful biochemical platform for combinatorial therapy, bioimaging, and biodiagnostics.     

    Reference

    Li, Fangfei; Lu, Jun; Liu, Jin; Liang, Chao; Wang, Maolin; Luyao; Li, Defang; Yao, Houzong;  Zhang, Qiulong; Wen, Jia; Zhang, Zong-Kang; Li, Jie; Lv, Quanxia; He, Xiaojuan; Guo, Baosheng; Guan, Daogang; Yu, Yuanyuan; Dang, Lei; Wu, Xiaohao; Li, Yongshu; Chen, Guofen; Jiang, Feng; Sun, Shiguo; Zhang, Bao-Ting; Lu, Aiping; Zhang, Ge (2017). A water-soluble nucleolin aptamer-paclitaxel conjugate for tumor-specific targeting in ovarian cancer. Nature Communications 1390, 8, 1.

     

    Zhang, X.-Q., Xu, X., Lam, R., Giljohann, D., Ho, D., & Mirkin, C. A. (2011). A Strategy for Increasing Drug Solubility and Efficacy through Covalent Attachment to Polyvalent DNA-Nanoparticle Conjugates. ACS Nano, 5(9), 6962–6970. http://doi.org/10.1021/nn201446c.

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  • 06/13/18--00:00: Methylene Blue Conjugates
  • The last two decades have seen an intensified development of nanoscale calorimetric and electrochemical sensors. Some of these sensors utilize on–target-inducible folding or unfolding of electrode-bound oligonucleotides which may also include aptamers or DNA switches. Examples are sensors for the detection of specific nucleic acids, proteins, small molecules, such as illicit drugs, and inorganic ions. Electrochemical DNA (E-DNA) and electrochemical aptamer-based sensors (E-AB) usually consist of an oligonucleotide probe modified with a redox reporter molecule at one terminal end.

    In most cases, the oligonucleotide conjugate is attached to a gold electrode via a thiol-gold bond at the other end. The redox reporter molecule used the most is methylene blue. In general, production for E-DNA and E-AB sensors can take 12 to 48 working hours.

    Methylene blue (Methylenblau) is a synthetic phenothiazine dye with redox properties first synthesized in 1876 by Heinrich Caro and tested by Paul Ehrlich in human patients. Ehrlich was able to demonstrate that it is useful for the treatment of malaria.

    Methylene blue (MB): C16H18ClN3S,  319.851 g/mol [Pubchem].

    “Methylene blue anhydrous is a compound consisting of dark green crystals or crystalline powder, having a bronze-like luster. Solutions in water or alcohol have a deep blue color. Methylene blue is used as a bacteriologic stain and as an indicator. It inhibits GUANYLATE CYCLASE, and has been used to treat cyanide poisoning and to lower levels of METHEMOGLOBIN.”  [Source: MeSH].

    “Methylene Blue is a synthetic basic dye. Methylene blue stains to negatively charged cell components like nucleic acids; when administered in the lymphatic bed of a tumor during oncologic surgery, methylene blue may stain lymph nodes draining from the tumor, thereby aiding in the visual localization of tumor sentinel lymph nodes. When administered intravenously in low doses, this agent may convert methemoglobin to hemoglobin.” [Source: NCIt].


    Also, methylene blue was found to be useful as a therapeutic dye having antimicrobial activity, allowing staining of living tissues, found to be useful in diagnostic histopathology, has blood staining activities, is used as a medicinal photosensitizer, for cancer chemotherapeutic uses and psychoactive uses in dementia and psychosis. Critical clinical applications of methylene blues include the therapy of methemoglobinemia, septic shock, encephalopathy, and ischemia. Methylene blue as a photosensitizer, a stain, and a redox indicator has many other applications as well. Methylene blue intercalates with the DNA duplex.

    Methylene blue’s autoxidizable chemical properties may be responsible for its unique biological action as both a metabolic energy enhancer and antioxidant that is frequently characterized by a biphasic dose-response.

    The blue colored oxidized form of methylene blue (MB) can be reduced to colorless Leuco-Methylene Blue (LMB). Methylene blue is an autoxidizing dye that at low concentrations is in equilibrium with leuko-methylene blue forming a reversible redox-oxidation system. Methylene blue in its oxidized form can accept electrons from an electron donor. The colorless reduced form, leuko-methylene blue, acts as an electron donor. The transfer of electrons to oxygen forms water. 

    Methylene blue is also used as a photosensitizer for the production of singlet oxygen or in photodynamic therapy for the treatment of cancer. Methylene Blue is widely used as a redox indicator in analytical chemistry. A redox indicator undergoes a color change at a specific electrode potential. 

    Methylene blue can be covalently attached to DNA using flexible C12 or Cn alkyl linker units. The final construct is a sensitive redox reporter for DNA based electrochemical measurements. As reported by Pheeney and Barton (2012), the reduction of methylene blue conjugated to a DNA duplex occurs by two mechanisms on a DNA-modified electrode:

    (1) Intercalated methylene blue tethered to the oligonucleotide duplex can be reduced in the DNA base pair stack, and 

    (2) by direct surface reduction at the electrode. 

    An example of an E-DNA sensor containing a stem-loop DNA modified on one terminal end with the redox reporter molecule and chemi-absorbed at the other end to a gold electrode is illustrated below.

    The hybridization of a complementary target DNA to the stem region brakes the stem and the beacon linearizes. The linearization of the oligonucleotide reduces the electron transfer efficiency and decreases the observed current. Regeneration of the sensor is possible by washing with ultrapure water at room temperature.

    Reference

    Buchholz, K., Schirmer, R. H., Eubel, J. K., Akoachere, M. B., Dandekar, T., Becker, K., & Gromer, S. (2008). Interactions of Methylene Blue with Human Disulfide Reductases and Their Orthologues from Plasmodium falciparum . Antimicrobial Agents and Chemotherapy, 52(1), 183–191. http://doi.org/10.1128/AAC.00773-07.

    Ferguson BS, Hoggarth DA, et al. (2013) Real-time, aptamer-based tracking of circulating therapeutic agents in living animals. Sci Transl Med, 5(213):213ra165.

    Garrido, E., Pla, L., LozanoTorres, B., ElSayed, S., MartínezMáñez, R., & Sancenón, F. (2018). Chromogenic and Fluorogenic Probes for the Detection of Illicit Drugs. ChemistryOpen, 7(5), 401–428. http://doi.org/10.1002/open.201800034.

    Liu Y, Tuleouva N, et al. (2010) Aptamer-based electrochemical biosensor for interferon gamma detection. Anal Chem, 82(19):8131–8136.

    Muren, N. B., & Barton, J. K. (2013). Electrochemical Assay for the Signal-on Detection of Human DNA Methyltransferase Activity. Journal of the American Chemical Society, 135(44), 16632–16640. http://doi.org/10.1021/ja4085918

    Oz, M., Lorke, D. E., Hasan, M., & Petroianu, G. A. (2011). Cellular and Molecular Actions of Methylene Blue in the Nervous System. Medicinal Research Reviews, 31(1), 93–117. http://doi.org/10.1002/med.20177

    http://www.glenresearch.com/GlenReports/GR30-15.html

    Sun, H., Zhu, X., Lu, P. Y., Rosato, R. R., Tan, W., & Zu, Y. (2014). Oligonucleotide Aptamers: New Tools for Targeted Cancer Therapy. Molecular Therapy. Nucleic Acids3(8), e182–. http://doi.org/10.1038/mtna.2014.32.

    Wainwright M, Crossley KB. Methylene Blue-a therapeutic dye for all seasons? J Chemother. 2002;14:431–443. [PubMed]

    Xiao Y, Lai RY, Plaxco KW. (2007) Preparation of electrode-immobilized, redox-modified oligonucleotides for electrochemical DNA and aptamer-based sensing. Nat Protoc, 2(11):2875–2880.

    Zhu, C., Yang, G., Li, H., Du, D., & Lin, Y. (2015). Electrochemical Sensors and Biosensors Based on Nanomaterials and Nanostructures. Analytical Chemistry, 87(1), 230–249. http://doi.org/10.1021/ac5039863

    ---...---


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    Minor groove binder (MGB) phosphoramidites are new tools that allow adding the MGB moiety dihydropyrroloindole-carboxylate (CDPI3) to the 3’- or 5’-end of oligonucleotides.

    MGB phosphoramidites can be used for the synthesis of modified oligonucleotides that contain the MGB moiety and have increased duplex stability useful in PCR, DNA array and antisense applications.


    Adding bridged nucleic acids to selected positions within the oligonucleotide sequence will increase the stability of the final DNA duplex even more.

    The structures of the two commercially CDPI3 MGB phosphoramiditesare shown below.

    Figure 1: 5’-CDPI3, MGB Phosphoramidite. This phosporamidite allows adding the MGB moiety to the 5’-end of an oligonucleotide during solid phase oligonucleotide synthesis.  

    Figure 2: CDPI3, MGB CPG. This phosporamidite allows adding the MGB moiety to the 3’-end of an oligonucleotide during solid phase oligonucleotide synthesis. 


    Kutyavin et al. in 2000 designed and synthesized a fluorogenic DNA probes conjugated to a minor groove binder moiety. These new MGB probes were more specific than standard DNA probes. The probes stabilized A/T rich duplexes more than G/C duplexes.

    Figure 3: DNA duplex formed with MGB-oligonucleotide probe.


    If a MGB-oligonucleotide probe is designed using FRET pairs such as a reported dye, for example, fluorescein, on the 5’-end, and an internal quencher, for example, TAMRA, upstream of the MGB moiety, the probe can be used for real-time measurements in PCR.

    The hybridized probe is cleaved during the primer extension step by the 5’-exonuclease activity of Taq polymerase which results in the release of a fluorescent signal. The fluorescent signal can be detected by any quantitative PCR instrument that combines thermal cycler functions with a fluorimeter.

    Several types of MGB probes have been developed and tested in recent years. These are:

    (1)        5’-F-3’-Q MGB TAQMAN probes

    (2)        3’-F-5’ Q FRET MGB probes

    (3)        3’-Q-5’-F FRET MGB probes

    (4)        miRNA inhibitor probes

    These new types of probes increase the repertoire of molecular tools useful for diagnostics and gene expression modulation.

    We at Biosynthesis Inc. have extensive experience in DNA and RNA synthesis as well as in probe design.


    Reference

    MGB-probes: http://www.glenresearch.com/GlenReports/GR29-11.html

    Kutyavin, I. V., Afonina, I. A., Mills, A., Gorn, V. V., Lukhtanov, E. A., Belousov, E. S., … Hedgpeth, J. (2000). 3′-Minor groove binder-DNA probes increase sequence specificity at PCR extension temperatures. Nucleic Acids Research, 28(2), 655–661. https://www.ncbi.nlm.nih.gov/pmc/articles/PMC102528/


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    Natural RNA tags added to RNA of interest allow detection, characterization, labeling and affinity purification of RNA target molecules. For example, small aptamer RNAs that bind with high affinity and specificity to Sephadex beads or streptavidin allow purification of intact RNA-protein complexes. Furthermore, RNA tags inserted into selected locations in genes encoding RNA components enable their detection. The CRISPR Cas system can be used together with synthetic RNA mimics to incorporate RNA tags into specific genomic locations to allow labeling.

    Recently a two-color CRISPR labeling system was developed by Wang and coworkers. Adding selective tags to a target-specific single guide RNA allows recruiting CRISPR-Cas systems for controlling gene expression.    

    Synthetic tagged RNA constructs allow purification of target RNAs and RNA complexes including miRNA and lncRNA as well as other molecules attached to the RNAs using pull-down techniques. RNA tags are unique tools for studying a variety of target RNAs including RNA transcripts, miRNAs, ncRNAs, lncRNAs, splicing event as well as others.

    Synthetic RNA can be chemically tagged or modified through the incorporation of modified ribonucleotides, such as BNAs, functional moieties such as biotin, fluorescent dyes, or natural or synthetic aptamers, for example, S1, D8, MS2 hairpin loops. Other types of RNA or constructs can be added as well.

    RNA affinity tags allow rapid enrichment of RNA binding protein (RNP) complexes from cellular lysates using mild conditions. This approach allows purification of RNP complexes under native conditions.

    Also, RNA tags can be used for the isolation of precursor or other RNA molecules of interest including RNPs.

    Known methods for the tagging of RNAs are: 

    (1)   Chemical tagging during in vitro transcription. 

    (2)   Incorporation of a well-characterized protein-binding RNA sequence
            during in vitro or in vivo transcription.

    (3)   Hybridization of affinity-tagged oligonucleotides that can also be biotinylated or
            modified, for example with BNAs.

    (4)   Incorporation of an artificially selected RNA motif during in vivo or
            in vitro transcription. 

    However, each of these methods has advantages and disadvantages.

    A variety of other RNA based applications are possible as well. Combining two or more hairpins within one RNA molecule is also possible.  

    Table 1:   Common RNA Tags

    Tag

    Sequence Info

    Notes

    Sephadex Tag

    D8 Sephadex RNA motif is recognized by Sephadex.

     

    5′ UCCGAGUAAU UUACGUUUUG  

       AUACGGUUGC GGA 3′

    The D8Sephadex-binding RNA minimal motif has 33 nucleotides. The indicated minimal structural motif has been discovered. The D8 tag was shown to bind specifically to Sephadex G-100 (Pharmacia).

    The D8 tag does not bind to other similar matrices such as Sepharose or Seph-acryl. binding of the tag to Sephadex can be efficiently competed with dextran B512.

    Steptavidin Tag

    S1 Streptavidin RNA motif recognized by streptavidin.

     

    5′ ACCGACCAGA AUCAUGCAAG

       UGCGUAAGAU AGUCGCGGGC

       CGGG 3′

     

    Kd of ~70 nM.

    The S1 streptavidin-binding RNA motif has 44 nucleotides originally selected to bind to streptavidin in either streptavidin–agarose bead assays or polyacrylamide gel electrophoretic mobility shift assays.

    Bound RNA tags can thus be released from streptavidin under otherwise native binding conditions by the inclusion of biotin in the binding buffer.

    MS2 Tag or

    MS2-TRAP

    Tagged RNA affinity purification.

    RNA is tagged with MS2 RNA hairpins and a fusion protein recognizing the MS2 RNA hairpins, MS2-GST is used.

    Identification of miRNAs associated with a target transcript in the cellular context.

    Identification of microRNAs that associate with a long intergenic (li)ncRNA.

    BoxB sequence

     = Boxb Rna

    NGTTCACCTCTAACCGGGTGAGCC

     

    Is recognized by the bacteriophage protein λ N:

     

    N Peptide: 

    ESKGTAKSRYKARRAELIAERR 

    BoxB can be used to tether proteins to RNAs, for example to mRNAs.

    PP7 Hairpin

    Binds to PP7 coat protein.

    Different RNA hairpins are rcognized by the coat proteins of different single-stranded RNA phages.

     

    Proposed and solved Structures of Hairpins


    Figure 1: Minimal binding motif and consensus structures of the Sephadex-affinity tag.

    Figure 2: Minimal binding motif and consensus structures of the streptavidin-affinity tag. (X indicates nonconserved nucleotides).

    Figure 3: Different views of the structural model of a MS2-RNA hairpin (G-5) complex. (Source: PDB 2C51). 

    Figure 4: Different views of the structural model of the MS2-RNA hairpin (G-5). (Source: PDB 2C51).

    pdb|1NYB|A Chain A, Solution Structure Of The Bacteriophage Phi21
               N Peptide-Boxb Rna Complex  ESKGTAKSRYKARRAELIAERR
    pdb|1NYB|B Chain B, Solution Structure Of The Bacteriophage Phi21
               N Peptide-Boxb Rna Complex  NGTTCACCTCTAACCGGGTGAGCC

    Figure 5: Two views of the structural models from the solution structure of a 22-amino-acid peptide from the amino-terminal domain of the bacteriophage φ21 N protein in complex with its cognate 24-mer boxB RNA hairpin solved with NMR.

    The nut (N utilization) site of bacteriophage lambda consists of two genetically defined elements, boxA and boxB. boxB forms an RNA hairpin and its 5 bp stem and 5 nt loop are recognized by the N peptide. The N peptide binds as an α-helix and interacts predominately with the major groove side of the 5′ half of the boxB RNA stem-loop. The φ21 boxB loop (CUAACC) has a structure typical of the “U-turn” motif.

    Figure 6: Two views of the PP7 coat protein dimer in complex with RNA hairpin. The RNA hairpin binds across the β-sheet surface of the coat protein dimer.

    The combined use of the MS2 hairpin with the PP7 hairpin allows detection of RNA in live cells if a chimeric protein consitent of the phage protein, a nuclear localization signal, and a fluorescent molecules is used. The MS2/PP7 appraoch has been used for the study of movement and localization of RNA as well the formation of RNA at the transcription site.    

    Reference

    Baron-Benhamou J, Gehring NH, Kulozik AE, Hentze MW.; Using the lambdaN peptide to tether proteins to RNAs. Methods Mol Biol. 2004;257:135-54. https://www.ncbi.nlm.nih.gov/pubmed/14770003
    Chao, J. A., Patskovsky, Y., Almo, S. C., & Singer, R. H. (2008). Structural basis for the coevolution of a viral RNA–protein complexNature Structural & Molecular Biology15(1), 103–105. 
    Cilley CD, Williamson JR.; Structural mimicry in the phage phi21 N peptide-boxB RNA complex. RNA. 2003 Jun;9(6):663-76. https://www.ncbi.nlm.nih.gov/pubmed/12756325

    Gesnel, M.-C., Del Gatto-Konczak, F., & Breathnach, R. (2009). Combined Use of MS2 and PP7 Coat Fusions Shows that TIA-1 Dominates hnRNP A1 for K-SAM Exon Splicing Control. Journal of Biomedicine and Biotechnology, 2009, 104853. http://doi.org/10.1155/2009/104853.

    Lenstra, T. L., & Larson, D. R. (2016). Single-Molecule mRNA Detection in Live YeastCurrent Protocols in Molecular Biology / Edited by Frederick M. Ausubel ... [et Al.]113, 14.24.1–14.24.15. http://doi.org/10.1002/0471142727.mb1424s113.

    Lo, A., & Qi, L. (2017). Genetic and epigenetic control of gene expression by CRISPR–Cas systems . F1000Research6, F1000 Faculty Rev–747. 

    Francis Lim and David S. Peabody; RNA recognition site of PP7 coat protein.Nucleic Acids Res. 2002 Oct 1; 30(19): 4138–4144. PMCID: PMC140551



    Marchese, D., de Groot, N. S., Lorenzo Gotor, N., Livi, C. M., & Tartaglia, G. G. (2016). Advances in the characterization of RNA
    binding proteins. Wiley Interdisciplinary Reviews. RNA, 7(6), 793–810. http://doi.org/10.1002/wrna.1378.

    Pascale Legault, Joyce Li, Jeremy Mogridge, Lewis E Kay, Jack Greenblatt; NMR Structure of the Bacteriophage λ N Peptide/boxB RNA Complex: Recognition of a GNRA Fold by an Arginine-Rich Motif. Cell, Volume 93, Issue 2, 1998, 289-299.https://doi.org/10.1016/S0092-8674(00)81579-2. (http://www.sciencedirect.com/science/article/pii/S0092867400815792)

    Srisawat, C., & Engelke, D. R. (2002). RNA affinity tags for purification of RNAs and ribonucleoprotein complexes. Methods (San Diego, Calif.), 26(2), 156–161. http://doi.org/10.1016/S1046-2023(02)00018-X

    Walker, S. C., Scott, F. H., Srisawat, C., & Engelke, D. R. (2008). RNA Affinity Tags for the Rapid Purification and Investigation of RNAs and RNA–Protein Complexes. Methods in Molecular Biology (Clifton, N.J.), 488, 23–40. http://doi.org/10.1007/978-1-60327-475-3_3

    Wang, S., Su, J.-H., Zhang, F., & Zhuang, X. (2016). An RNA-aptamer-based two-color CRISPR labeling system. Scientific Reports, 6, 26857. http://doi.org/10.1038/srep26857.

    Yoon, J.-H., Srikantan, S., & Gorospe, M. (2012). MS2-TRAP (MS2-tagged RNA affinity purification): Tagging RNA to identify associated miRNAs. Methods (San Diego, Calif.), 58(2), 81–87. http://doi.org/10.1016/j.ymeth.2012.07.004

    Yoon, J.-H., & Gorospe, M. (2016). Identification of mRNA-interacting factors by MS2-TRAP (MS2-tagged RNA affinity purification). Methods in Molecular Biology (Clifton, N.J.), 1421, 15–22. http://doi.org/10.1007/978-1-4939-3591-8_2

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    Bioinformatic algorithms allowing predicting of biomolecular folding for proteins, peptides, and RNAs, even though sometimes successful, have all their limitations. The reasons are as follows.

    •   RNA molecules in solution may adopt secondary structures that are only partially
         determined by thermodynamics since RNA molecules can undergo conformational
         changes during interaction with other RNAs, RNA binding proteins or RNA binding
         peptides. These interactions are very complex and difficult to model.

    •   Our knowledge of thermodynamic rules and parameters that govern folding
         patterns of RNAs are far from being complete.

    •   Most folding algorithms use approximations for scanning the landscape of possible
         secondary structures. Therefore the predicted structures are often just an
         estimate or approximation and chemical methods are needed to verify the
         deduced structures.

    •   Predicting pseudoknots and long-range and tertiary-structure interactions
         accurately is also quite difficult.

    •   Using experimental methods to accumulate additional experimental data
         should allow improvements in algorithms used in computational methods.    

    The mapping of RNA-protein or RNA-RNA interactions by protein pull-down or affinity pull-down methods allow studying RNA structures, as well as RNA-protein, and RNA-RNA interactions. Since RNA-binding proteins (RBPs) are key players in the post-transcriptional regulation of gene expression precise knowledge of their binding sites is critical for determining their molecular function and for understanding their roles in cell development and disease.

    NMR

    Nuclear magnetic resonance spectroscopy is a powerful tool for studying RNA structures in detail. NMR allows studying RNA molecules in a more natural state when dissolved in solution. The drawback is that a large preparation of highly purified and uniform RNA is needed, and it is limited to solving small structures. 
    https://www.ncbi.nlm.nih.gov/pubmed/14523911,  https://www.nature.com/articles/ncomms8024.


    RIP

    RNA immunoprecipitation (RIP) uses antibodies to pull down RNA bound to a targeted protein. RIP allows detection of individual proteins associated with specific nucleic acid regions on RNA molecules. Live cells are treated with formaldehyde to generate protein-RNA cross-links between molecules that are nearby. Cross-linked RNA connected to the targeted protein is isolated using immunoprecipitation of the protein. Recovery and quantitative analysis of the immunoprecipitated RNAs are achieved by reversal of the formaldehyde cross-linking which now permits reverse transcription using PCR. RIP is similar to chromatin immunoprecipitation (ChiP). Unfortunately, the technique cannot differentiate between direct and indirectly bound RNA. A drawback of this method is that the generation of false positives from interaction occurring after cell lysis is also possible. 
    https://www.ncbi.nlm.nih.gov/pubmed/27659976


    CLIP

    Cross-linking and immunoprecipitation (CLIP) allows mapping of transcriptome-wide binding sites of RNA-binding proteins. RNA-protein complexes are covalently cross-linked and purified from intact tissue cells. CLIP improved the specificity of RIP. It allows the removal of weekly bound RNA when stringent washing steps are used. RNAs that remain can be reversed transcribed, and PCR amplification allows sequencing with next-generation sequencing. For CLIP to work, reverse transcription needs to proceed from a universal 3′ ligated adapter to a universal 5′ ligated adapter. Both adapters are required for PCR amplification.

    All CLIP protocols use RNA-protein cross-linking and immunoprecipitation targeting a protein of interest.  Ultraviolet light creates the cross-links between RNA and proteins in vivo. The RNA is then isolated and reverse transcribed into cDNA. The cDNA can be used on several platforms to identify and quantify interacting RNAs. Several refinements and specializations of this central CLIP principle exist: CLIP-seq, PAR-CLIP, and iCLIP are three of the most common.

    http://epigenie.com/quick-review-crosslinking-immunoprecipitation-clip-methods/
    https://www.ncbi.nlm.nih.gov/pubmed/3071669
    http://science.sciencemag.org/content/302/5648/1212.long
    https://www.ncbi.nlm.nih.gov/pubmed/16314267?dopt=Abstract
    https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4492529/https://en.wikipedia.org/wiki/CLIP


    iCLIP

    The individual-nucleotide resolution CLIP (iCLIP) protocol was developed to allow recovery of truncated cDNAs lost in CLIP. iCLIP enables PCR amplification of truncated cDNAs and identifies protein–RNA crosslink sites with single nucleotide resolution. iCLIP identifies protein-RNA cross-links on a genome-wide scale. An intramolecular cDNA circularization step enables analysis of cDNAs truncated at the protein-RNA crosslink sites with high resolution and specificity. A 3’ exonuclease degrades protein-bound RNA. The enzyme digests the isolated RNA but stops at the cross-linked protein. An adapter is then ligated to the remaining RNA.

    For iCLIP, cells are irradiated with UV-C light on ice. Covalent bonds are formed between proteins and RNA. The cross-linking reaction is followed by partial RNase digestion and immunoprecipitation with protein-specific antibodies. Libraries are preparation and visualized by dephosphorylation of RNA. Next, a 3′ end adapter is ligated to the RNA, and the 5′ end is radioactively labeled. Complexes are separated by SDS–PAGE and isolated from a nitrocellulose membrane. Proteins are digested by proteinase K, and reverse transcription (RT) is performed truncating at the remaining polypeptide. An RT primer introduces two cleavable adapter regions and barcode sequences. Free RT primers are removed by size selection, and circularization of the cDNA is carried out. Linearization generates suitable templates for PCR amplification.Finally, high-throughput sequencing generates sequencing reads in which a barcode sequence is immediately followed by the last nucleotide of the cDNA.
    https://www.ncbi.nlm.nih.gov/pubmed/24184352 
    https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3988997/
    http://epigenie.com/quick-review-crosslinking-immunoprecipitation-clip-methods/


    eCLIP

    Enhanced CLIP improves library preparation and circular steps of iCLIP. eCLIP simplifies the generation of paired IgG and size-matched input controls. Also, eCLIP improves specificity in the discovery of authentic binding sites. After dephosphorylation of RNA fragments, an “inline barcoded” RNA adapter is ligated to the 3′ end. Following protein gel electrophoresis and nitrocellulose membrane transfer, a region of 75 kDa (~220 nt of RNA) above the protein size is excised and proteinase K treated to isolate RNA. The RNA is further prepared into paired-end high-throughput sequencing libraries and sequenced.
    https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4887338/

    PAR-CLIP

    Photo-activable ribonucleoside-enhanced cross-linking and immunoprecipitation (PAR-CLIP) is a method that allows identification of binding sites of RNA-binding proteins (RBPs). PAR-CLIP can be used for the identification of RNA-protein binding sites in transcriptomes. For the method to work, photoreactive ribonucleoside analogs are incorporated into nascent RNA transcripts in living cells. The use of ultraviolet (UV) light of 365 nm cross-links photoreactive nucleoside-labeled cellular RNAs to RNA-binding proteins.

    First, photo-reactive thioribonucleosides are incorporated into nascent transcripts. Cross-linking is achieved by irritating the cells with ultraviolet long-wavelengths greater than 310 nm (365 nm is usually used).  Immunoprecipitation is used for the purification of cross-linked RNA–RBP complexes which are further purified by SDS-PAGE. The recovered RNA is converted into a cDNA library and is sequenced using next-generation sequencing. Multiple sequencing platforms can be used.

    Reverse transcription of cross-linked RNA with incorporated photoactivable thioribonucleosides, followed by PCR amplification, results in a characteristic mutation (T-to-C when using 4SU and G-to-A when using 6SG) that is used to identify the RNA recognition elements. 
    https://www.ncbi.nlm.nih.gov/pubmed/29236251


    DMS Treatment

    Dimethylsulfate (DMS, C2H6O4S, 126.13 g/mol) treatment modifies RNA by adding a methyl group to any unpaired or loosely structured adenosine (A) and cytidine (C) in oligonucleotides. DMS mapping is one of the oldest chemical RNA mapping methods. After methylation, the bases can no longer form base pairs and will cause cDNA transcripts to terminate early allowing identification of the presence of unpaired bases. The following use of next-generation-sequencing (DMS-seq/Structure-seq) increases the power of the technique and allows quantitative mapping of base modifications. Targeted Structure-seq further improves the specificity of the technique by using primers targeting the length of a specific RNA of interest without having to sequence the whole transcriptome. 
    https://www.ncbi.nlm.nih.gov/pmc/articles/PMC2701642/


    PARS

    Parallel analysis of RNA structure (PARS) combines classical RNA foot-printing methods with next-generation sequencing. Here, poly(A) selected RNA is folded in vitro and incubated with either RNase V1 or S1 nuclease to probe for double- and single-stranded regions. RNase V1 and S1 nuclease cleavage results in a 5′P leaving group. Next, the RNA is fragmented. Enzymatic cleavage products contain 5′P, but fragmentation and degradation products have 5′OH groups. Only true structure-probing sites can be ligated to adaptors and reverse transcribed. Sequencing the resulting cDNA library using high-throughput sequencing and mapping the resulting reads to the genome allow identification of double- or single-stranded regions in the transcriptome. Different PARS scores indicate that a base is either double-stranded (positive score) or single-stranded (negative score).


    PARIS

    Psoralen analysis of RNA interactions and structure (PARIS) is a psoraleb based cross-linking method that works by fixing base pairs of double-stranded RNA (dsRNA) of cells in vivo using the specific cross-linker 4′-aminomethyl-trioxsalen (AMT). PARIS analysis directly determines transcriptome-wide base pairing interactions. PARIS combines in vivo cross-linking, 2D gel purification, proximity ligation, and high-throughput sequencing to allow high-throughput and near-base pair resolution determination of the RNA structures and interactions in living cells. After cross-linking sample are treated with proteinase and the RNA is partially degradated. The resulting sequencing reads represent all native dsRNAs in the organism. These can be mapped to infer their structure.
    https://www.ncbi.nlm.nih.gov/pubmed/29130190


    Psoralen conjugated oligonucleotides

    Psoralen conjugated oligonucleotides can also be used as antisense oligonucleotides (ASOs). Triple helix-forming oligonucleotides linked to psoralen (pso-TFO) allow introduction of DNA interstrand cross-links at specific sites in the genome of living mammalian cells. 
    Co-introduction of duplex DNA with target region homology results in precise knock-in of the donor at frequencies of 2 to 3 orders.

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    The development of next-generation sequencing (NGS) methods revolutionized the study of genomic transcriptionally active regions. NGS now makes it possible to catalog transcripts from coding sequences and also allows the discovery of RNA species that are not templates for protein synthesis.

    Several classes of non-coding RNAs (ncRNAs), including microRNAs (miRNAs) and long non-coding RNAs (ncRNAs), are already known to play diverse roles in post-transcriptional regulation of mRNA stability and epigenetic control of chromatin activities.

    Recently a new class of RNAs called enhancer RNAs or eRNAs has been identified {Kim et al. 2010}. However, it remains an open question whether eRNAs are just transcriptional noise or have a relevant biologically function. As usually, to clarify this, more research is needed.

    Figure 1: Overview of a typical gene transcribed by RNA polymerase II.

    A typical gene transcribed by RNA polymerase II has a promotor that extends upstream from the site where transcription is initiated (Figure 1).  (See Lewin, Benjamin; Genes VII chapter 20, 2000). The promotor contains several short sequence elements that bind transcription factors, up to 10 base pairs (bp) in length, and promotors are dispersed over a sequence region that can be greater than 200 bp. Enhancers contain a more closely packed array of elements that also bind transcription factors. Enhancer regions may be located at a distance of several kilobases (kb). The DNA duplex may be coiled or rearranged such that transcription factors at the promotor and the enhancer interact to form a large protein-DNA complex.

    What are eRNAs?

    Enhancer RNA or eRNA are RNAs transcribed by RNA polymerase II (RNAPII) from the domain of transcription enhancers. eRNAs are a class of short non-coding RNAs, 50 to 2,000 nucleotides in length, transcribed from DNA enhancer regions. eRNAs stimulate gene expression, but the precise mechanisms how they function remains unclear. Enhancers are intergenic DNA elements that regulate transcription of target genes in response to signaling pathways by interacting with promotors over large genomic distances. Enhancers contain binding sites for transcription factors that promote RNA polymerase II (RNAPII) recruitment and transcription activation. Also, enhancers carry unique epigenetic marks that distinguish them from promoters. Additionally, these regulatory elements have an open chromatin conformation that increases accessibility to transcription factors and RNAPII. Enhancers are bi-directionally transcribed into eRNAs {Kim et al. 2010}.

    Rahman et al. recently reported that eRNAs are localized exclusively in the nucleus and the induction of eRNAs occurs with similar kinetics as that of target mRNAs. eRNAs are mostly nascent at enhancers however their steady-state levels remain lower than those of their cognate mRNAs at the single-allele level. eRNAs are rarely co-expressed with their target loci. It appears that active gene transcription does not require the continuous transcription of eRNAs or their accumulation at enhancers. 

    Genome-wide sequencing methods allowed studying stimulus-dependent enhancer functions in tissues cells. Kim et al. in 2010 found that the level of eRNA expression at neuronal enhancers positively correlates with the level of mRNA synthesis in nearby genes. Their findings suggest that eRNA synthesis occurs specifically at enhancers that actively promote mRNA synthesis and the mechanism of enhancer activation involves RNAPII binding and eRNA synthesis.

    Levels of eRNA expression correlate with mRNA levels of the corresponding enhancers target gene. eRNAs can be identified by a specific chromatin signature: H3K4me1 and H3K27ac. Furthermore, enhancers are often transcribed and bound by RNA polymerase II. eRNAs appear to be rarely spliced, are not polyadenylated, and are often transcribed in both directions. So far, the mechanisms by which eRNAs facilitate enhancer function remain unclear. More research to clarify this is needed.

    Kim et al. suggested that eRNA synthesis is required to establish and maintain a chromatin landscape at enhancers needed for enhancer function. It is also possible that the eRNA transcripts are functionally important by themselves.

    Enhancers are classified as cis-regulatory genetic elements that controll temporal and cell-type specific patterns of gene expression. Active enhancers have been found to generate bi-directional non-coding RNA transcripts called enhancer eRNAs. Enhancers contain bidirectional elements that allow assisting initiation and the activity of a promotor is increased by the presence of an enhancer. The enhancer is located distinct from the promotor and the position of the enhancer relative to the promotor can vary substantially. An enhancer can stimulate any promotor placed in its vicinity. Enhancers often show redundancy in function. DNA must be able to form a loop structure if proteins bound at an enhancer several kb distant from a promotor need to interact directly with proteins bound in the vicinity of the starting point such that the enhancer and promotor are closely located to each other. Enhancers may function by bringing proteins into the vicinity of the promotor. A model for eRNAs and transcriptional activation of a typical gene is illustrated in figure 2.

    Figure 2: Model for eRNAs, enhancer derived RNAs, and transcriptional activation of a gene (Feng Liu, 2017). The structure of a typical gene is illustrated. The gene is associated with two cis-regulatory elements, (i) a promotor, located in proximity, and (ii) an enhancer, located distal to the transcription start site of the gene. In general, when its enhancer is inactivae the gene is turned “off” (A). When an enhancer is activated by transcription factors, it can loop towards the promotor and turn “on” the transcription of the gene (B). Both, enhancers and promotors are classified as non-coding elements.



    A more detailed model can be reviewed in the paper published by Kim et al. in 2015 in which the loop structure is shown together with the transcription machinery.

    To find and varify the presence of eRNAs, Espinosa suggested in 2015 to design a couple of CRISPRs, one to delete the lncRNA "promotor" and a second one to induce a pol(A) cassette early in the path of RNAPII. If the first genome edit affects expression but the second one doesn't it is an "e" or eRNA. 



    Reference

    Espinosa JM; Revisiting lncRNAs: How Do You Know Yours Is Not an eRNA? Mol Cell. 2016 Apr 7;62(1):1-2. doi: 10.1016/j.molcel.2016.03.022.

    Giles et al. 2015. ncRNA function in chromatin organization., in Epigenetic Gene Expression and Regulation}.

    Kim, T.-K., Hemberg, M., Gray, J. M., Costa, A. M., Bear, D. M., Wu, J., … Greenberg, M. E. (2010).
    Widespread transcription at neuronal activity-regulated enhancers. Nature, 465(7295), 182–187. http://doi.org/10.1038/nature09033. https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE21161.

    Kim, T-K., Hemberg, M., and Gray J.M.; 2015. Enhancer RNAs: A class of long noncoding RNAs synthesized at enhancers. Cold Spring Harb. Perspect 2015,7:a018622, 1-3. 

    Lewin, Benjamin; Genes VII chapter 20, 2000

    Feng Liu; Enhancer-derived RNA: A Primer. Genomics Proteomics Bioinformatics 15 (2017) 196-200.

    Rahman, S., Zorca, C. E., Traboulsi, T., Noutahi, E., Krause, M. R., Mader, S., & Zenklusen, D. (2017).
    Single-cell profiling reveals that eRNA accumulation at enhancer–promoter loops is not required to sustain transcription. Nucleic Acids Research, 45(6), 3017–3030. http://doi.org/10.1093/nar/gkw1220.

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    BNAs dramatically improve the specificity of CRISPR-Cas9. 

    The addition of BNAs (2′,4′-BNANC[N-Me]) to CRISPR-RNAs (crRNAs) dramatically improves the accuracy in CRISPR-based gene-editing. In this approach, natural crRNA molecules are replaced with synthetic crRNAs containing bridged nucleic acids, or BNAs.

    BNA-T (2′,4′-BNANC[N-Me]-T)

    Off-target effects or off-target cutting and the generation of additional mutations remain a significant barrier for the use of Cas9-based gene editing. Therefore a number of approaches have been investigated and developed to improve the specificity of Cas9 and to minimize or abolish off-target effects. 

    Cromwell et al. in 2018 now report that the incorporation of bridged nucleic acids, BNAs, at specific positions within crRNAs significantly improves Cas9 DNA cleavage specificity in vitro and cells with close to no off-target cleavage. The researcher reasoned that modified crRNAs enhance specificity by impairing the formation of the stable “zipped” conformation during hybridization to off-target sequences.

    These results indicate that BNA modified crRNAs improve the specificity of the CRISPR-Cas9 system and illustrate the power of recently developed synthetic nucleic acid technologies to solve problems in enzyme specificity as well.

    The research was published in "Nature Communications" in a paper entitled “Incorporation of Bridged Nucleic Acids into CRISPR RNAs Improves Cas9 Endonuclease Specificity.”

    The CRISPR (Clustered Regularly Interspaced Short Palindromic Repeat)-Cas9 complex is a component of prokaryotic immune systems that is now a widely used tool for genome editing technologies. CRISPR-Cas9 can be used for the generation of gene knock-outs and knock-ins in a variety of organisms, functional genomics studies, epigenetic screens, and proof-of-principle studies that enable the correction of genetic diseases in mammals.

    Reference

    Cromwell, Christopher R., Sung, Keewon, Park, Jinho, Krysler, Amanda R., Jovel, Juan, Kim, Seong Keun, and Hubbard, Basil P.; 2018. Incorporation of bridged nucleic acids into CRISPR RNAs improves Cas9 endonuclease specificity. Nature Communications 9, 1448.


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    Horseradish Peroxidase, or HRP, allows the design and synthesis of molecular probes such as in situ hybridization probes to increase signal amplification. HRP, when conjugated to hybridization probes, increases signal amplification, for example, in Catalyzed Reporter Deposition-Fluorescent In Situ Hybridization (CARD-FISH) applications.

    Historically, horseradish peroxidase is used as a reporter enzyme in diagnostics and histochemistry applications. Horseradish Peroxidase can be conjugated to 
    oligonucleotides, peptides, proteins, or other molecules, for example, via a maleimide attachment to form a thioester bond.


    Horseradish Peroxidase is an enzymatically active glycoprotein. HRP is a globular molecule with an α-helical secondary structure and one short β-sheet region (Gajhede et al. 1997). 

    The HRP molecule is separated into a distal and a proximal region by an iron protoporphyrin IX cofactor. The iron protoporphyrin IX cofactor is known as the heme group. Heme is typically linked to HRP by a coordinate bond of the heme iron with a conserved His170 residue. In the presence of H2O2, HRP catalyzes an oxidation reaction that converts reduced, non-fluorescent or non-colorimetric substrates into oxidized, fluorescent or colorimetric molecules. Furthermore, HRP C1A contains nine Asn-X-Ser/Thr-X motives (X being any amino acid but Pro) which are potential N-glycosylation sites.

    For conjugation reactions, surface lysine residues allow directed enzyme immobilization or conjugation to target compounds via covalent linkages. On the surface of HRP C1A, three out of six lysine residues were found accessible to chemical modifications. These are 
    Lys174, Lys232, and Lys241 (O’Brien et al. 2001). 


    Reference

    Ferrari, B. C., Tujula, N., Stoner, K., & Kjelleberg, S. (2006). Catalyzed Reporter Deposition-Fluorescence In Situ Hybridization Allows for Enrichment-Independent Detection of Microcolony-Forming Soil Bacteria. Applied and Environmental Microbiology, 72(1), 918–922. [PubMed]

    Gajhede M, Schuller DJ, Henriksen A, Smith AT, Poulos TL. Crystal structure of horseradish peroxidase C at 2.15 A resolution. Nat Struct Biol. 1997;4:1032–8. [
    PubMed]

    Krainer, F. W., & Glieder, A. (2015). An updated view on horseradish peroxidases: recombinant production and biotechnological applications. Applied Microbiology and Biotechnology, 99, 1611–1625. 

    [PubMed]

    O’Brien AM, O’Fágáin C, Nielsen PF, Welinder KG. Location of crosslinks in chemically stabilized horseradish peroxidase: implications for the design of crosslinks. Biotechnol Bioeng. 2001;76:277–84. [
    PubMed]


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    To increase solubility and effectiveness of Paclitaxel, the conjugation of the drug to gold nanoparticles via DNA linkers is possible.  According to Zhang et al., resulting conjugates usually show a 50-fold increase in solubility in aqueous buffers in comparison to the unconjugated drug. For visualization of the conjugation reaction and the effect of the drug conjugate in cells, the DNA linker can be labeled with fluorophores as well (Zhang et al. 2011). The final nano-gold DNA Paclitaxel conjugate represents a multimodal drug delivery system with

    (i)  enhanced solubility of the drug in aqueous systems, and serum-containing
         cell culture media,

    (ii) increased drug efficacy in Paclitaxel-resistant cells, and

    (iii) a useful way of tracking the delivery of the drug. 

    However, other types of Paclitaxel conjugates are also possible.



    Taxol

    .

    Paclitaxel, Taxol A,  (Source: Drugbank)

    (2AR-(2aalpha,4beta, 4abeta,6beta, 9alpha(alpha r*,betas*), 11alpha,12alpha,12balpha))-beta-(benzoylamino)-alpha- hydroxy-benzenepropanoic acid 6,12b-bis(acetyloxy)-12-(benzoyl-oxy)-2a,3,4,4a,5,6,9,10,11,12,12a,12b-dodecahydro-4,11-dihydroxy-4a,8,13,13-tetramethyl-5-oxo-7,11-methano-1H-cyclodeca(3,4)benz(1,2-b)oxet-9-yl ester. 

    C47H51NO14.

    Mw: Average: 853.9061 
    ; Monoisotopic: 853.330955345


    “Paclitaxel is a mitotic inhibitor used in cancer chemotherapy. It was discovered in a US National Cancer Institute program at the Research Triangle Institute in 1967 when Monroe E. Wall and Mansukh C. Wani isolated it from the bark of the Pacific yew tree, Taxus brevifolia and named it taxol. Later it was discovered that endophytic fungi in the bark synthesize paclitaxel. When it was developed commercially by Bristol-Myers Squibb (BMS), the generic name was changed to paclitaxel and the BMS compound is sold under the trademark Taxol. In this formulation, paclitaxel is dissolved in Kolliphor EL and ethanol, as a delivery agent. A newer formulation, in which paclitaxel is bound to albumin, is sold under the trademark Abraxane”.  [Wikipedia, Drugbank]
     

    Taxol binds to microtubules in the cell and slows down cell division and growth. It does so by stabilizing microtubules thereby blocking the segregation of chromosomes resulting in cell death or cellular apoptosis. However, formulation of Taxol into a delivery system acceptable for human use proved difficult. According to the NIH, Taxol is the best-selling drug ever manufactured.

    Different illustrations of the tubulin structure with highlighted GTP (non-exchangeable) bound to α-tubulin, GDP (exchangeable) bound to β-tubulin, and Taxol.




     

    Due to its inherent insolubility in aqueous media, Paclitaxel can be limited in effectiveness. However, the covalent attachment to gold nanoparticles via DNA linkers results in conjugates highly soluble in aqueous buffers. Zhang et al. in 2011 reported a 50 fold increase in solubility of the conjugates when compared to the unconjugated drug.

    The research group recommends the attachment of DNA-nanoparticle conjugates to drugs as a strategy for the enhancement of a wide variety of therapeutic agents that need to be formulated in aqueous media.  


    Reference

    Hermanson, Grg T.; Bioconjugate Techniques.

    Kim CK, Ghosh P, Rotello VM.;  Multimodal drug delivery using gold nanoparticles. Nanoscale. 2009 Oct;1(1):61-7. doi: 10.1039/b9nr00112c. Epub 2009 Sep 4.

    Zhang, X.-Q., Xu, X., Lam, R., Giljohann, D., Ho, D., & Mirkin, C. A. (2011). A Strategy for Increasing Drug Solubility and Efficacy through Covalent Attachment to Polyvalent DNA-Nanoparticle Conjugates. ACS Nano, 5(9), 6962–6970. http://doi.org/10.1021/nn201446c

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    To overcome disadvantages of drugs with poor water solubility different strategies such as the conjugation of the drug to small molecules, carbohydrates, oligonucleotides, peptides, or other molecules have been developed by many research groups from all over the world in recent decades.

    The solubility of the drug Paclitaxel in aqueous media can be increase by conjugating Paclitaxel to aptamers, oligonucleotides, or DNA linked gold nanoparticles.

    A good example is the conjugation of functionalized Paclitaxel to gold nanoparticles via a DNA linker. In this approach Zhang et al. used succinic anhydride to add a functional extender group to the drug that now allows conjugation to oligonucleotides containing an amino group on one of it's terminal ends followed by conjugating this conjugate to nano-gold particles. First, Paclitaxel is reacted with succinic anhydride to generated Paclitaxel carboxylic acid which now allows conjugation to an amino group on a DNA oligonucleotide.

    The following steps were needed for the production of fluorescent Paclitaxel-DNA-gold nanoparticles.


    (1)    Synthesize propyl thiol-(3’-end) DNA oligomers with a terminal amino group
             (5’-end) needed for covalent attachment of Paclitaxel.

    (2)    Modify Paclitaxel with succinic anhydride to create Paclitaxel carboxylic acid.

    (3)    Covalently attach Paclitaxel carboxylic acid to the modified oligonucleotide
             using EDC/Sulf-NHS.

    (4)    Purify and QC the derivative (PTX-DNA) using MALDI-MS.

    (5)    Immobilize the derivative on citrate-stabilized gold nanoparticles (AuNPs).

    (6)    Remove excess PTX-DNA.

    (7)    Resuspend PTX-DNA-AuNPs.

    For the synthesis of the Paclitaxel-fluorescein-DNA conjugate (Zhang et al. 2011) succinic anhydride was used to modify Paclitaxel by the reaction with succinic anhydride to create Paclitaxel carboxylic acid. 




    A modified DNA oligonucleotide containing a amino group at the 5’-end, an internal fluorophore, and a thiol group at the 3'-end was covalently attached to Paclitaxel carboxylic acid to yield the Paclitaxel-DNA (PTX-DNA) compound. (Structural models are not to scale).


    Immobilization of PTX-DNA on citrate-stabilized gold nanoparticles (AuNPs) yielded the Paclitaxel-DNA gold nanoparticle conjugate (PTX-DNA@AuNPs). HPLC is used for the purification of intermediate products and MALDI-TOF-MS is used for QA/QC. 



    Zhang et al. showed that covalently attached hydrophobic Paclitaxel onto gold nanoparticles via a DNA linker resulted in enhanced hydrophilicity and stability of the conjugate. Cellular internalization, delivery, and distribution of the drug conjugate was monitored in human breast adenocarcinoma cells and uterine sarcoma cells with the help of confocal fluorescence microscopy. Furthermore, the cell-killing activity of paclitaxel was enhanced in vitro against several cancer cell lines when delivered as the gold nanoparticle conjugate.

    The authors of the referenced papers suggested that this approch and other similar approaches can be used for other biological useful compounds such as peptides, small interfering RNA (siRNA), gadolinium complexes, antibodies, as well as aptamers.

    To conclude, these types of conjugates represent a powerful biochemical platform for combinatorial therapy, bioimaging, and biodiagnostics.     

    Reference

    Li, Fangfei; Lu, Jun; Liu, Jin; Liang, Chao; Wang, Maolin; Luyao; Li, Defang; Yao, Houzong;  Zhang, Qiulong; Wen, Jia; Zhang, Zong-Kang; Li, Jie; Lv, Quanxia; He, Xiaojuan; Guo, Baosheng; Guan, Daogang; Yu, Yuanyuan; Dang, Lei; Wu, Xiaohao; Li, Yongshu; Chen, Guofen; Jiang, Feng; Sun, Shiguo; Zhang, Bao-Ting; Lu, Aiping; Zhang, Ge (2017). A water-soluble nucleolin aptamer-paclitaxel conjugate for tumor-specific targeting in ovarian cancer. Nature Communications 1390, 8, 1.

     

    Zhang, X.-Q., Xu, X., Lam, R., Giljohann, D., Ho, D., & Mirkin, C. A. (2011). A Strategy for Increasing Drug Solubility and Efficacy through Covalent Attachment to Polyvalent DNA-Nanoparticle Conjugates. ACS Nano, 5(9), 6962–6970. http://doi.org/10.1021/nn201446c.

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  • 06/13/18--00:00: Methylene Blue Conjugates
  • The last two decades have seen an intensified development of nanoscale calorimetric and electrochemical sensors. Some of these sensors utilize on–target-inducible folding or unfolding of electrode-bound oligonucleotides which may also include aptamers or DNA switches. Examples are sensors for the detection of specific nucleic acids, proteins, small molecules, such as illicit drugs, and inorganic ions. Electrochemical DNA (E-DNA) and electrochemical aptamer-based sensors (E-AB) usually consist of an oligonucleotide probe modified with a redox reporter molecule at one terminal end.

    In most cases, the oligonucleotide conjugate is attached to a gold electrode via a thiol-gold bond at the other end. The redox reporter molecule used the most is methylene blue. In general, production for E-DNA and E-AB sensors can take 12 to 48 working hours.

    Methylene blue (Methylenblau) is a synthetic phenothiazine dye with redox properties first synthesized in 1876 by Heinrich Caro and tested by Paul Ehrlich in human patients. Ehrlich was able to demonstrate that it is useful for the treatment of malaria.

    Methylene blue (MB): C16H18ClN3S,  319.851 g/mol [Pubchem].

    “Methylene blue anhydrous is a compound consisting of dark green crystals or crystalline powder, having a bronze-like luster. Solutions in water or alcohol have a deep blue color. Methylene blue is used as a bacteriologic stain and as an indicator. It inhibits GUANYLATE CYCLASE, and has been used to treat cyanide poisoning and to lower levels of METHEMOGLOBIN.”  [Source: MeSH].

    “Methylene Blue is a synthetic basic dye. Methylene blue stains to negatively charged cell components like nucleic acids; when administered in the lymphatic bed of a tumor during oncologic surgery, methylene blue may stain lymph nodes draining from the tumor, thereby aiding in the visual localization of tumor sentinel lymph nodes. When administered intravenously in low doses, this agent may convert methemoglobin to hemoglobin.” [Source: NCIt].


    Also, methylene blue was found to be useful as a therapeutic dye having antimicrobial activity, allowing staining of living tissues, found to be useful in diagnostic histopathology, has blood staining activities, is used as a medicinal photosensitizer, for cancer chemotherapeutic uses and psychoactive uses in dementia and psychosis. Critical clinical applications of methylene blues include the therapy of methemoglobinemia, septic shock, encephalopathy, and ischemia. Methylene blue as a photosensitizer, a stain, and a redox indicator has many other applications as well. Methylene blue intercalates with the DNA duplex.

    Methylene blue’s autoxidizable chemical properties may be responsible for its unique biological action as both a metabolic energy enhancer and antioxidant that is frequently characterized by a biphasic dose-response.

    The blue colored oxidized form of methylene blue (MB) can be reduced to colorless Leuco-Methylene Blue (LMB). Methylene blue is an autoxidizing dye that at low concentrations is in equilibrium with leuko-methylene blue forming a reversible redox-oxidation system. Methylene blue in its oxidized form can accept electrons from an electron donor. The colorless reduced form, leuko-methylene blue, acts as an electron donor. The transfer of electrons to oxygen forms water. 

    Methylene blue is also used as a photosensitizer for the production of singlet oxygen or in photodynamic therapy for the treatment of cancer. Methylene Blue is widely used as a redox indicator in analytical chemistry. A redox indicator undergoes a color change at a specific electrode potential. 

    Methylene blue can be covalently attached to DNA using flexible C12 or Cn alkyl linker units. The final construct is a sensitive redox reporter for DNA based electrochemical measurements. As reported by Pheeney and Barton (2012), the reduction of methylene blue conjugated to a DNA duplex occurs by two mechanisms on a DNA-modified electrode:

    (1) Intercalated methylene blue tethered to the oligonucleotide duplex can be reduced in the DNA base pair stack, and 

    (2) by direct surface reduction at the electrode. 

    An example of an E-DNA sensor containing a stem-loop DNA modified on one terminal end with the redox reporter molecule and chemi-absorbed at the other end to a gold electrode is illustrated below.

    The hybridization of a complementary target DNA to the stem region brakes the stem and the beacon linearizes. The linearization of the oligonucleotide reduces the electron transfer efficiency and decreases the observed current. Regeneration of the sensor is possible by washing with ultrapure water at room temperature.

    Reference

    Buchholz, K., Schirmer, R. H., Eubel, J. K., Akoachere, M. B., Dandekar, T., Becker, K., & Gromer, S. (2008). Interactions of Methylene Blue with Human Disulfide Reductases and Their Orthologues from Plasmodium falciparum . Antimicrobial Agents and Chemotherapy, 52(1), 183–191. http://doi.org/10.1128/AAC.00773-07.

    Ferguson BS, Hoggarth DA, et al. (2013) Real-time, aptamer-based tracking of circulating therapeutic agents in living animals. Sci Transl Med, 5(213):213ra165.

    Garrido, E., Pla, L., LozanoTorres, B., ElSayed, S., MartínezMáñez, R., & Sancenón, F. (2018). Chromogenic and Fluorogenic Probes for the Detection of Illicit Drugs. ChemistryOpen, 7(5), 401–428. http://doi.org/10.1002/open.201800034.

    Liu Y, Tuleouva N, et al. (2010) Aptamer-based electrochemical biosensor for interferon gamma detection. Anal Chem, 82(19):8131–8136.

    Muren, N. B., & Barton, J. K. (2013). Electrochemical Assay for the Signal-on Detection of Human DNA Methyltransferase Activity. Journal of the American Chemical Society, 135(44), 16632–16640. http://doi.org/10.1021/ja4085918

    Oz, M., Lorke, D. E., Hasan, M., & Petroianu, G. A. (2011). Cellular and Molecular Actions of Methylene Blue in the Nervous System. Medicinal Research Reviews, 31(1), 93–117. http://doi.org/10.1002/med.20177

    http://www.glenresearch.com/GlenReports/GR30-15.html

    Sun, H., Zhu, X., Lu, P. Y., Rosato, R. R., Tan, W., & Zu, Y. (2014). Oligonucleotide Aptamers: New Tools for Targeted Cancer Therapy. Molecular Therapy. Nucleic Acids3(8), e182–. http://doi.org/10.1038/mtna.2014.32.

    Wainwright M, Crossley KB. Methylene Blue-a therapeutic dye for all seasons? J Chemother. 2002;14:431–443. [PubMed]

    Xiao Y, Lai RY, Plaxco KW. (2007) Preparation of electrode-immobilized, redox-modified oligonucleotides for electrochemical DNA and aptamer-based sensing. Nat Protoc, 2(11):2875–2880.

    Zhu, C., Yang, G., Li, H., Du, D., & Lin, Y. (2015). Electrochemical Sensors and Biosensors Based on Nanomaterials and Nanostructures. Analytical Chemistry, 87(1), 230–249. http://doi.org/10.1021/ac5039863

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    Pseudouridine (), one of the most abundant modified RNA molecules, is synthesized in situ by  synthases. Naturally occurring RNA molecules are often modified. Over 100 different types of RNA modifications have been identified and were known for decades, however recent advances in analytical technologies have revealed that they appear to function in nearly every class of cellular RNA. It is now becoming apparent that in both coding and noncoding RNAs, dynamic modifications add a new layer of control of genetic information. Many years ago sequencing of the first biological RNA (1965) identified ten modifications including pseudouridine () in the alanine transfer RNA from yeast. 

    Oligonucleotides modified with pseudouridine are useful tools to investigate how pseudouridine modified RNAs function in vivo and in vitro. Pseudouridine modified oligonucleotides can be synthesized using pseudouridine phosphoramidites and are available at Biosynthesis Inc.

    Figure 1:  Structure of the ring-open pseudo-uridine mono-phosphate (ΨMP) from the crystal structure of ΨMP glycosidase/ ring-opened ribose ΨMP complex obtained by co-crystallizing ΨMP glycosidase with 4 mM MnCl2, 2 mM R5P and saturated uracil (PBD ID: 4GIL. Huang et al. 2012).

     

    Modifications are derived from the standard nucleosides, adenosine, guanosine, cytidine and uridine. RNA modifications are present in many different types of RNA in the cell. RNA modifications are generated by enzymatic processing of the corresponding primary transcript. A large number of enzymes and pathways catalyzing post-transcriptional RNA modifications are known.   

    Figure 2:  Conversion of uridine to pseudouridine.

    Pseudouridine is synthesized by conversion of uridine. The reaction involves breaking the bond that connects the ribose to the N1 of uridine and reconnecting the ribose to position C5 of the uridine. Uridine synthases are proteins known to function without RNA cofactors. 
    Pseudouridine is the result of isomerization or internal trans-glycosylation of uridine (U) into pseudouridine (), catalyzed by RNA:pseudouridine synthases. The reaction is also called a base-specific isomerization.

    Pseudouridine derived from isomerization of the uridine base is the most common modification in cellular RNA and abundantly found in ribosomal RNA (rRNA) and transfer RNA (tRNA). Pseudouridine is present in ~ 1 to 2 % of “salt-soluble” RNA fractions (tRNA) and < 1% in “salt-insoluble” fractions (rRNA). Recently, Pseudouridine-sequencing (PseudU-seq) established the presence of in human and yeast mRNAs as well. A chemical labeling and pull-down method (CeU-seq) identified over 2,000 sites in human mRNA.

    Li et al. in 2015 showed with the help of quantitative mass spectrometry that Ψ is much more prevalent (Ψ/U ratio ∼0.2-0.6%) in mammalian mRNA than previously believed. The research group developed N3-N-cyclohexyl-N’-b-(4-methyl-morpholinium) ethylcarbodiimide (CMC)-enriched pseudouridine sequencing (CeU-Seq), a selective chemical labeling and pulldown method, to identify 2,084 Ψ sites within 1,929 human transcripts, of which four (in ribosomal RNA and EEF1A1 mRNA) were biochemically verified.

    Ψ modifications are dynamically regulated by the pseudouridine synthase (Pus) enzymes. Pus enzymes are known to catalyze the isomerization of Ψ in response to stress conditions such as heat shock. Ψ also affects the secondary structure of RNA. In addition, Ψ functions in altering stop codon read through which may also be biologically relevant.

    Reference

    Grosjean, H.; Modification and editing of RNA: historical overview and important facts to remember. In Topics in Current Genetics pp 1-22. In: Grosjean H. (eds) Fine-Tuning of RNA Functions by Modification and Editing. Topics in Current Genetics, vol 12. Springer, Berlin, Heidelberg.

    Huang, S., Mahanta, N., Begley, T. P., & Ealick, S. E. (2012). Pseudouridine Monophosphate Glycosidase: a New Glycosidase Mechanism,. Biochemistry, 51(45), 9245–9255. http://doi.org/10.1021/bi3006829https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3526674/

    Lewin, B.; Genes VII, Oxford University Press. 2000, pp. 715.



    Li, X., Zhu, P., Ma, S., Song, J., Bai, J., Sun, F., and Yi, C.
    Chemical pulldown reveals dynamic pseudouridylation of the mammalian transcriptome. Nat. Chem. Biol.2015; 11: 592–597. https://www.ncbi.nlm.nih.gov/pubmed/26075521?dopt=Abstract



    Roundtree, Ian A. et al.; Dynamic RNA Modifications in Gene Expression Regulation.Cell , Volume 169 , Issue 7 , 1187 – 1200.

    Yi, C., & Pan, T. (2011). Cellular Dynamics of RNA Modification. Accounts of Chemical Research, 44(12), 1380–1388. http://doi.org/10.1021/ar200057m. https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3179539/



    Zaringhalam M
    , Papavasiliou FN.;  Pseudouridylation meets next-generation sequencing. Methods. 2016 Sep 1;107:63-72. doi: 10.1016/j.ymeth.2016.03.001. Epub 2016 Mar 8.



    Zhao, B. S., & He, C. (2015). Pseudouridine in a new era of RNA modifications. Cell Research, 25(2), 153–154.
    http://doi.org/10.1038/cr.2014.143.

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    The nucleoside analogue 3-cyanovinyl carbazole, CNVK, enables ultrafast reversible photo-cross-linking of oligonucleotides.

    Recently a new non-enzymatic and non-invasive technologies for nucleic acid editing has been reported that takes advantage of the photo-cross-linker 3-cyanovinyl carbazole (CNVK). 3-Cyanovinyl carbazole (CNVK) when incorporated into oligonucleotides photo-cross-links a target pyrimidine and the CNVK to afford deamination of cytosine converting it to uracil.

    Figure 1: 3-Cyanovinyl carbazole (CNVK) nucleoside analog. This nucleoside is now available as a cyanoethyl (CE) phosphoramidite allowing the incorporation of the analog into oligonucleotides during automated solid phase oligonucleotide synthesis.


    Cross-linking and photo-cross-linking reactions are used widely in biochemistry, biology, chemistry, and molecular biology for the study of molecular interactions. Many new cross-linking reagents have been developed and synthesized in the last two decades. Advances made in analytical instrumentation now allow the analysis of cross-linked products at ever lower levels of detection. For example, various photo-cross-linking approaches can be used to study the interactome of cells.

    The interactome refers to the network of molecular interactions in living cells, such as protein-protein, protein-nucleic acid, or protein-carbohydrate interactions.

    Photo-cross-linking is used fo the study of protein-protein or protein-oligonucleotide interactions in living cells. In this approach, first photo-cross-linkers are incorporated in the desired proteins or oligonucleotides within cells, next, cells are exposed to UV radiation to induce the cross-linking between molecules in contact to the cross-linking functional groups. Following cross-linking, the target molecule of interest and its cross-linked complexes are isolated using immune-purification and analyzed using mass spectrometry. Often enzymatic digestions are used to allow for easier detection of the cross-linked products and to define molecular interphases.


    The new photo-cross-linker 3-cyanovinylcarbazole (

    CNVK) 

    nucleoside allows the study of oligonucleotide interactions.

    Yoshimura and Fujimoto in 2008 descibed a ultrafast reversible photo-cross-linking reaction for the manipulation of DNA in situ using CNVK which was applied to the selection of RNA in 2009 by the same research group.

    When CNVK is incorporated into oligonucleotides, a very rapid cross-link is formed to a pyrimidine base on the complementary strand when irridated at 366 nm.

    According to Yoshimura and Fujimoto, photo-cross-linking to thymine takes only 1 second but takes 25 second to cytosine.

    Complete reversal of the cross-link is achieved by irridation at 312 nm for 3 minutes. 
     




    Figure 2: Photo-cross-linking reaction of the 3-Cyanovinyl carbazole (CNVK) oligonucleotide with a thymidine on a complementary oligonucleotide strand. The photoadduct CNVK-T is shown as the reaction product as described by Yohimura and Fujimoto in 2008 and 2009



    Figure 2: Photo-cross-linking reaction of the 3-Cyanovinyl carbazole (CNVK) oligonucleotide with a cytosine at 366 nm. Heading the cross-link to 90 oC for 3.5 hours causes deamination of the cytosine to form a uracil in the complementary strand (Yoshimura et al. 2009). Reversal of the cross-link at 312 nm results in a complementary strand with a uracil in place of the cytosine. 
     

    In
    2010 

    Fujimoto et al. reported the cross-linking reaction of CNVK incorporated into oligonucleotides to a dC residue in duplex DNA. The heating of the cross-link to 90 °C for 3.5 hours resulted in a deamination reaction of the cytosine base now forming a uracil moiety in the complementary strand. Reversal of the cross-link at 312 nm resulted in a DNA strand in which dC is replaced with dU.  The transformation is specific for a dC residue opposite a CNVK residue. However, any further adjacent dC residues are unaffected by the reaction. Also, oligonucleotides containing CNVK residues can be cross-linked to adjacent RNA strands. 

    In 2013, this approach was applied to SNP-based genotyping and in 2015 a threoninol linker was introduced in the oligonucleotide strand.


    Sethi et al. in 2018 reported that the ultrafast reversible DNA inter-strand photo-cross-linking reaction of oligodeoxynucleotides (ODNs) containing CNVK can take place at physiological conditions. in order to find the best combination of photo-active nucleobase and complementary base of the target, a variety of combinatios were studied.
    The study concluded that 

    the best combination of counter-base and photo-cross-linker will  result in the highest conversion of C→U and in the least time at physiological conditions.

    Reference

    2008.   Y. Yoshimura, and K. Fujimoto; Ultrafast reversible photo-cross-linking reaction: toward in situ DNA manipulation. Org Lett. 2008 Aug 7;10(15):3227-30. doi: 10.1021/ol801112j. Epub 2008 Jun 27.

    2009.   Y. Yoshimura, T. Ohtake, H. Okada, and K. Fujimoto; A new approach for reversible RNA photocrosslinking reaction: application to sequence-specific RNA selection. ChemBioChem, 2009, 10, 1473-6.

    2010. Fujimoto K1, Konishi-Hiratsuka K, Sakamoto T, Yoshimura Y.; Site-specific cytosine to uracil transition by using reversible DNA photo-crosslinking.Chembiochem. 2010 Aug 16;11(12):1661-4. doi: 10.1002/cbic.201000274.

    2012. Wong and Jameson; Chemistry of protein and Nucleic Acid Cross-Linking and Conjugation. 2nd Edition. CRC Press. 2012.

    2013.   Kenzo Fujimoto, Asuka Yamada, Yoshinaga Yoshimura, Tadashi Tsukaguchi, and Takashi Sakamoto; Details of the Ultrafast DNA Photo-Cross-Linking Reaction of 3-Cyanovinylcarbazole Nucleoside: Cis–Trans Isomeric Effect and the Application for SNP-Based Genotyping. J. Am. Chem. Soc., 2013, 135 (43), pp 16161–16167.  DOI: 10.1021/ja406965f. Publication Date (Web): October 2, 2013. Copyright © 2013 American Chemical Society kenzo@jaist.ac.jp  :J. Am. Chem. Soc.  135, 43, 16161-16167

    2013. Pham, N. D., Parker, R. B., & Kohler, J. J. (2013). Photocrosslinking approaches to interactome mapping. Current Opinion in Chemical Biology, 17(1), 90–101. http://doi.org/10.1016/j.cbpa.2012.10.034. 

    2015Sakamoto T, Tanaka Y, Fujimoto K.; DNA photo-cross-linking using 3-cyanovinylcarbazole modified oligonucleotide with threoninol linker.   Org Lett. 2015 Feb 20;17(4):936-9. doi: 10.1021/acs.orglett.5b00035. Epub 2015 Feb 5. 

    2018Sethi, S.; Nakamura, S.; Fujimoto, K.; Study of Photochemical Cytosine to Uracil Transition via Ultrafast Photo-Cross-Linking Using Vinylcarbazole Derivatives in Duplex DNA. Molecules 2018, 23, 828.

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    Photocleavable oligonucleotides have many applications in biological and medicinal research. Photoactive groups can function as protecting groups of oligonucleotides and for a variety of other bioactive or reporter molecules. Molecules functionalized with a photoactive group are also known as caged compounds. Photocleavable or photolabile groups have been used heavily in synthetic organic chemistry for a variety of applications. For example, molecular releasing systems utilize photochemical reactions based on energy transfer between a quencher or sensitizer and the photoactive group. Even molecular logic gates or networks can be designed using multiple components in combination with photocleavable groups or linker molecules.

    The o-nitrobenzyl derivatives have gained enormous popularity in synthetic chemistry and as building blocks for oligonucleotide and peptide synthesis.

    The function of the photolabile group is based on the photoisomerization of an o-nitrobenzyl alcohol derivative into a corresponding o-nitrobenzaldehyde upon irradiation with UV light and the release of a free carboxylic acid (Zhao et al. 2012: o-nitrobenzyl alcohol derivatives).



    Figure 1:  Structure of the PC-Spacer Phosphoramidite [4-(4,4'-Dimethoxytrityloxy) butyramidomethyl)-1-(2-nitrophenyl)-ethyl]- 2-cyanoethyl-(N,N-diisopropyl)-phosphoramidite].



    Figure 2: Example of oligonucleotide attachment points to the PC spacer moiety. This non-nucleosidic photocleavable spacer molecule can be incorporated into oligonucleotides either on the 5 ‘- or 3’-end. If only incorporated on the 5’-end oligonucleotides can be immobilized on glass slides or other surfaces. This spacer type is used in a variety of applications, for example, for the synthesis of photocleavable block copolymers. To test if the photocleavage is quantitative a fluorophore can be attached at one end such that the dye is released in a way that allows monitoring of the reaction. For example, from 5’- tagged oligonucleotides immobilized to a surface on the 3’-end. 


    Figure 3: A photocleavable biotin spacer useful for phosphoramidite oligonucleotide synthesis.  This spacer allows the capture of modified oligonucleotides using the streptavidin system and the rapid, quantitative removal of the biotin spacer using near UV light at 300 to 350 nm. The released oligonucleotides are now suitable for other biological applications such as gene construction as part of a cloning experiment.   

    Applications of 5’- Photocleavable Oligonucleotides:

    Affinity isolation and purification of nucleic acids binding proteins

    Cassette mutagenesis.

    Diagnostic assays requiring release of the probe-target complex or specific markers.

    DNA-programmed single oxygen production.

    DNAzyme deactivation and reactivation.

    Multiple non-radioactive probes for DNA/RNA blots.

    Nucleic acid sensing.

    Nucleobase-Caging.

    Optochemical control of cellular processes.

    Optochemical control of DNA:Protein interactions.

    PCR.

    Photoactivated molecular beacons for drug release.

    Photosensitized Singlet Oxygen Production.

    Spatially-addressable photorelease of probe-target complexes or marker molecules for diagnostics.

    Templated catalysis.

    Examples from the literature:

    Olejnik and others in 1998 reported the synthesis of oligonucleotides containing a photocleavable amino group conjugated to the 5'-terminal phosphate of oligonucleotides. This 5' photocleavable amino group allows the introduction of various amine-reactive molecules to synthetic oligonucleotides as well as their immobilization on activated solid supports. Irradiation with near-UV light selectively cleaves off the photocleavable bond on the 5'-phosphate. Photocleavable conjugates containing biotin, digoxigenin and fluorophores such as tetramethylrhodamine can be prepared as well.

    Clo and others in 2006 demonstrated an on-and-off switching 1O2 sensitizer controlled by DNA sequences. The photosensitizer pyropheophorbide-a (P) is attached or conjugated to a short 15-mer nucleotide sequence and a “black hole quencher 3” (Q) is attached to a 21-mer oligonucleotide which complements the 15-mer oligonucleotide. P and Q are brought into close proximity after DNA assembly by hybridization. The singled state of P is quenched by Q via FRET or by contact-mediated electron exchange.

    Zhou et al. in 2012 described a mass spectrometry-based assay for the analysis of protein kinase activities in a multiplexing format. This assay can be used for screening inhibitors against multiple kinases in parallel useful for drug discovery and predictive diagnostics.

    For this assay to work, oligonucleotides are tagged or conjugated to peptides. The peptides selected are substrates for kinases allowing efficient capture from solution-phase kinase reactions by annealing to the complementary sequence tethered to PEG-passivated superparamagnetic microparticles.

    Synthesis of peptide-oligonucleotide conjugates with bifunctional photocleavable cross-linkers enables a reversible conjugation. The relative state of phosphorylation is detected using mass spectrometry after washing away contaminants and photoreleasing the peptides.

    Reference

    Emiliano Cló, John W. Snyder, Niels V. Voigt, Peter R. Ogilby, and, and Kurt V. Gothelf; DNA-Programmed Control of Photosensitized Singlet Oxygen Production. Journal of the American Chemical Society2006128 (13), 4200-4201. DOI: 10.1021/ja058713a.  https://pubs.acs.org/doi/abs/10.1021/ja058713a.

    Olejnik, J., Krzymanska-Olejnik, E., & Rothschild, K. J. (1998). Photocleavable aminotag phosphoramidites for 5’-termini DNA/RNA labeling. Nucleic Acids Research26(15), 3572–3576.  https://academic.oup.com/nar/article/26/15/3572/2360254.

    Zhou, G., Khan, F., Dai, Q., Sylvester, J. E., & Kron, S. J. (2012). Photocleavable Peptide-Oligonucleotide Conjugates for Protein Kinase Assays by MALDI-TOF MS. Molecular bioSystems, 8(9), 2395–2404. http://doi.org/10.1039/c2mb25163a.

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